Water quality monitoring protocols for streams and - PA DEP

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OFFICE OF WATER PROGRAMS

BUREAU OF CLEAN WATER

WATER QUALITY MONITORING PROTOCOLS FOR STREAMS AND RIVERS

2021

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Prepared by: Dustin Shull and Josh Lookenbill PA Department of Environmental Protection Office of Water Programs Bureau of Clean Water 11th Floor: Rachel Carson State Office Building Harrisburg, PA 17105 2018

Edited by: Josh Lookenbill and Rebecca Whiteash PA Department of Environmental Protection Office of Water Programs Bureau of Clean Water 11th Floor: Rachel Carson State Office Building Harrisburg, PA 17105

Cover image created by:

Matthew Shank, PA Department of Environmental Protection

Cover photographs by:

Top, bottom center, and bottom right: Matthew Shank, PA Department of Environmental

Protection

Bottom left: Chris Custer, Pennsylvania State University

2021

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TABLE OF CONTENTS

Table of Contents ............................................................................................................ ii

Abbreviations .................................................................................................................. v

Introduction .................................................................................................. 1-1

Purpose..................................................................................................................... 1-2

Updates..................................................................................................................... 1-4

Collection Framework and Processes ....................................................................... 1-4

Sampling Design and Planning .................................................................... 2-1

Types of Sampling Design ........................................................................................ 2-3

Sampling Plans ......................................................................................................... 2-4

2.2 Cause and Effect Surveys .................................................................................. 2-7

2.3 Combined Sewer Overflow (CSO) Compliance Data Collection ...................... 2-15

Biological Data Collection Protocols ............................................................ 3-1

3.1 Wadeable Riffle-Run Stream Macroinvertebrate Data Collection Protocol ........ 3-2

3.2 Wadeable Limestone Stream Macroinvertebrate Data Collection Protocol ........ 3-9

3.3 Wadeable Multihabitat Stream Macroinvertebrate Data Collection Protocol .... 3-14

3.4 Semi-Wadeable Large River Macroinvertebrate Data Collection Protocol ....... 3-20

3.5 Qualitative Benthic Macroinvertebrate Data Collection Protocol ...................... 3-28

3.6 Macroinvertebrate Laboratory Subsampling and Identification Protocol .......... 3-33

3.7 Fish Data Collection Protocol ........................................................................... 3-44

3.8 Fish Tissue Data Collection Protocol ............................................................... 3-64

3.9 Mussel Data Collection Protocol ...................................................................... 3-74

3.10 Periphyton Data Collection Protocol .............................................................. 3-88

3.11 Bacteriological Data Collection Protocol ...................................................... 3-143

Chemical Data Collection Protocols ............................................................ 4-1

4.1 In-Situ Field Meter and Transect Data Collection Protocol ................................. 4-2

4.2 Discrete Water Chemistry Data Collection Protocol ........................................... 4-9

4.3 Continuous Physicochemical Data Collection Protocol .................................... 4-25

4.4 Sediment Chemistry Data Collection Protocol ................................................. 4-93

4.5 Passive Water Chemistry Data Collection Protocol ....................................... 4-111

4.6 Sample Information System (SIS) Data Entry Protocol .................................. 4-120

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Physical Data Collection Protocols .............................................................. 5-1

5.1 Stream Habitat Data Collection Protocol ............................................................ 5-2

5.2 Pebble Count Data Collection Protocol .............................................................. 5-8

5.3 Water Flow Data Collection Protocol ............................................................... 5-14

Appendix A: Biological Data Collection Information and Forms ................................... A-1

A.1 Macroinvertebrate Field Data Sheet .................................................................. A-2

A.2 Macroinvertebrate Enumeration Bench Sheet ................................................... A-5

A.3 Fish Field Data Collection Sheet ....................................................................... A-7

A.4 Fish Tissue Field Data Collection Sheet .......................................................... A-15

A.5 Mussel Field Data Collection Sheets ............................................................... A-18

A.6 QBE Periphyton Field Data Collection Sheet ................................................... A-21

A.7 QMH Periphyton Field Data Collection Sheet .................................................. A-24

A.8 Algae Identification and Enumeration Laboratory Submission Data Sheet ...... A-27

A.9 Microbiology Sample Submission Sheet for Acidified Chlorophyll-a,

Phycocyanin, and AFDM ........................................................................................ A-29

A.10 Bacteria Monitoring Field and Site Data Sheet .............................................. A-31

Appendix B: Chemical Data Collection Information and Forms .................................... B-1

B.1 Field Meter Calibration Form ............................................................................. B-2

B.2 Transect Data Collection Form .......................................................................... B-4

B.3 Sample Information System User’s Guide ......................................................... B-7

B.4 Sediment Field Data Collection Sheet ............................................................. B-43

B.5 CIM Deployment Form ..................................................................................... B-46

B.6 CIM Maintenance Visit Field Form ................................................................... B-49

B.7 Temperature Adjustments for pH Buffers ........................................................ B-54

B.8 DO 100% Saturation Chart for Fresh Water .................................................... B-57

Appendix C: Physical Data Collection Information and Forms .................................... C-1

C.1 Riffle/Run Prevalence Habitat Data Collection Form ........................................ C-2

C.2 Riffle/Run Prevalence Habitat Data Collection Abbreviated (One-Page)

Form ........................................................................................................................ C-5

C.3 Low Gradient Habitat Data Collection Form ..................................................... C-7

C.4 Pebble Count Data Collection Form ............................................................... C-10

C.5 Alternative Pebble Count Data Collection Form ............................................. C-12

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Appendix D: Pennsylvania’s Surface Water Qulaity Monitoring Network (WQN)

Manual ........................................................................................................................ D-1

D.1 Standard Analysis Codes (SAC) .................................................................... D-14

D.2 Active Water Chemistry Stations .................................................................... D-19

D.3 Active Benthic Macroinvertebrate Stations ..................................................... D-31

D.4 Discontinued Stations ..................................................................................... D-44

D.5 WQN Objectives ............................................................................................. D-58

D.6 Field Preparation and Fixative Chart .............................................................. D-64

D.7 Stream Flow Measurement Protocols ............................................................. D-67

Appendix E: Quality Assurance Project Plan (QAPP): Data Collection for the Purpose

of Existing Use Evluations, Protected Use Assessment, and Evaluations of Point and

Nonpoint Source Impacts to Surface Water ................................................................. E-1

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ABBREVIATIONS

AFDM Ash Free Dry Mass

AIS Aquatic Invasive Species

AL Auditing Laboratory

ALU Aquatic Life Use

ANS Academy of Natural Sciences

AWS Wildlife Water Supply

BAH Best Available Habitat

BAT Best Available Technology

BCD Buoyancy Control Device

BOD Biological Oxygen Demand

BOL Bureau of Laboratories

CBOD Carbonaceous Biological Oxygen Demand

CIM Continuous Instream Monitoring

CPOM Coarse Particulate Organic Matter

CWF Cold Water Fishes

DELTP Deformities, Erosions, Lesions, Tumors and Parasites

DEP Pennsylvania Department of Environmental Protection

DCNR Pennsylvania Department of Conservation and Natural Resources

DES PFBC Division of Environmental Services

DO Dissolved Oxygen

DOH Pennsylvania Department of Health

DTH Depositional Targeted Habitat

DWM Diving Safety Manual

DWQ Division of Water Quality

EDC Endocrine Disrupting Compounds

EPT Ephemeroptera, Plecoptera, and Trichoptera

EST Environmental Sampling Technologies

EV Exceptional Value Waters

FNU Formazin Nephelometric Units

GPS Global Positioning System

GRTS Generalized Random Tessellation Stratified

HDPE High-Density Polyethylene

HQ High Quality Waters

HUC Hydrologic Unit Code

IBI Index of Biotic Integrity

ICE Instream Comprehensive Evaluation

IRS Irrigation Water Supply

IWS Industrial Water Supply

LDB Left Descending Bank

LIS Line-Intercept Sampling

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LWS Livestock Water Supply

MF Migratory Fishes

MMI Multi-Metric Index

MSDS Material Safety Data Sheets

NAWQA National Water-Quality Assessment

NELAP National Environmental Laboratory Accreditation Program

NOAA National Oceanic and Atmospheric Administration

NPDES National Pollutant Discharge Elimination System

NHD National Hydrologic Dataset

NTU Nephelometric Turbidity Units

PAH Polycyclic Aromatic Hydrocarbons

PCB Polychlorinated Biphenyls

PEP Pennsylvania Epilithic Periphyton Sampler

PFBC Pennsylvania Fish and Boat Commission

PFD Personal Floatation Device

PHY Phytoplankton

PL Participating Laboratory

POCIS Polar Organic Chemical Integrative Sampler

PPE Personal Protective Equipment

PRC Performance Reference Compounds

PWS Potable Water Supply

QA Quality Assurance

QBE Quantitative Benthic Epilithic

QC Quality Control

QMH Qualitative Multi-Habitat

qPCR Quantitative Polymerase Chain Reaction

RBP Rapid Bioassessment Protocol

RDB Right Descending Bank

RTH Richest Targeted Habitat

SAC Standard Analysis Code

SAV Submerged Aquatic Vegetation

SCP Scientific Collectors Permit

SIS Sample Information System

SOP Standard Operating Procedure

SPMD Semi-Permeable Membrane Device

SRBC Susquehanna River Basin Commission

TDS Total Dissolved Solids

TE Threatened and Endangered Species

TL Total Length

TMDL Total Maximum Daily Loads

TSF Trout Stocking

UDO Unit Diving Officer

USEPA United States Environmental Protection Agency

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USGS United States Geological Survey

VOA Volatile Organic Analysis

WQD Water Quality Division

WQN Water Quality Network

WQS Water Quality Standards

WWF Warm Water Fishes

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Prepared by:

Josh Lookenbill and Dustin Shull

PA Department of Environmental Protection

Office of Water Programs

Bureau of Clean Water

11th Floor: Rachel Carson State Office Building

Harrisburg, PA 17105

2017

Edited by:

Josh Lookenbill




11th Floor: Rachael Carson State Office Building


2021

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PURPOSE

This document details the suite of monitoring and data collection protocols currently

used by the Pennsylvania Department of Environmental Protection (DEP) to meet

multiple surface water characterization objectives in flowing waterbodies. This book is

part of a larger conceptual framework (Figure 1) that DEP uses to collect quality data

and make decisions on various surface water matters across Pennsylvania. Data

collection protocols have been compiled in this book entitled Water Quality Monitoring

Protocols for Streams and Rivers (Monitoring Book) to increase operational

transparency and to facilitate the use of DEP data collection protocols. The assessment

of protected water uses is one of the primary monitoring objectives of DEP staff

biologists, however there are several additional and equally important monitoring

objectives described throughout this document. At this time, lake and reservoir data

collection protocols are not included in this document, but DEP intends on updating and

adding them in the future.

Figure 1. Conceptual framework that DEP uses to make scientifically defensible data

collections and decisions.

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UPDATES

This is the first update to the Monitoring Book since the initial 2018 publication that

effectively consolidated DEP data collection protocols for flowing waterbodies. Data

collection protocols for lakes have not been consolidated into this document. This 2021

update includes the following updates that are worth noting:

• A new subchapter has been added to Chapter 2, Sampling Design and Planning

(Lookenbill 2020b), titled Combined Sewer Overflow (CSO) Compliance

Monitoring (Lookenbill 2020a). This new subchapter was developed in response

to requests by DEP permitting staff for guidance in reviewing post-construction

monitoring plans. The updates are primarily design and planning elements that

rely on existing data collection protocols.

• Updates to the Taxonomic Identification section of the Macroinvertebrate

Laboratory Subsampling and Identification Protocol (Brickner 2020) located in

Chapter 3, Biological Data Collection Protocols were made that more definitively

describe the appropriate levels of macroinvertebrate identification and the

process for addressing less than confident identifications.

• Updates to the Bacteriological Data Collection Protocol (Miller 2021) located in

Chapter 3 were made to account for the adoption of updated bacteriological

criteria in 25 Pa. § Code 93.7 during the swimming season (May 1 through

September 30).

• New sections were added to the In-Situ Field Meter and Transect Data Collection

Protocol (Hoger 2020) found in Chapter 4, titled Evaluation of Transect Data and

Display and Storage of Transect Data.

• Appendix D has been added to include the Water Quality Network (WQN). This

new appendix, Pennsylvania’s Surface Water Quality Network (WQN) Manual

(Pulket and Lookenbill 2020), was once a separate document and has now been

incorporated into this Monitoring Book to take advantage of existing resources

and consolidate this information.

• Appendix E has been added to include the latest EPA approved Quality

Assurance Protocol (QAPP).

COLLECTION FRAMEWORK AND PROCESSES

Collectors will need to adhere to an established framework and process. In doing so,

consistency will be maintained to produce quality data. All data collection should follow

the framework described here and throughout this book.

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• IDENTIFY THE MONITORING OBJECTIVE – Why are data being collected?

The primary and any secondary objectives will need to be identified before any

data are collected. DEP staff will often identify the primary data collection

objective and consider other, secondary objectives which may require additional

data collections, which would simultaneously meet requirements of both the

primary and secondary objectives. Consequently, considering both objectives

would require less effort if pursued independently. As an example, DEP staff

biologists will identify the primary objective as the assessment of protected water

uses. Based on the information gathered during the reconnaissance process,

there may be indications that the target waterbody may have an existing use that

is different than the designated use. Employing additional requirements that

satisfy the stream redesignation evaluation objective requirements, like the

collection of reference stations and targeting additional analytical tests, could

provide the opportunity to meet this secondary objective. The following is a list of

monitoring objectives:

o Assessment of Protected Water Uses

o Stream Redesignation Evaluation

o Point of First Use

o Cause and Effect

o Water Quality Network Monitoring

o Protocol, Method and Water Quality Standards Development

• RECONNAISANCE AND DATA GATHERING – Reconnaissance includes the

delineation of the targeted waterbody or basin and the gathering of historical and

other information that will provide the best opportunity to meet data collection

objectives.

• SELECT A SAMPLING DESIGN – Select a sampling design that will meet the

monitoring objective(s).

o Probabilistic

o Targeted

• DEVELOP A SAMPLING PLAN – A sampling plan should be developed that

outlines objectives, protocols, methods, standards, and other important

information and resources that need to be referenced and followed.

• QUALITY ASSURANCE – Water quality data that will be used by DEP will need

to meet quality assurance (QA) criteria. A review of the QA criteria for specific

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objectives will need to be performed before any collection occurs. Water quality

data that are used by DEP need to be collected by individuals trained and

subsequently audited by DEP staff. Documentation of trainings and audits will

need to be provided upon request. DEP maintains documentation of trainings

and audits as part of the QA requirements.

Monitoring for The Assessment of Protected Water Uses

DEP staff biologists are tasked with monitoring and assessing surface waters of

Pennsylvania for the protected water uses listed at 25 Pa. Code § 93.3. Monitoring to

assess protected water uses is one of the primary objectives of DEP staff biologists. To

successfully complete an assessment of protected water uses, requirements should be

met as described in the appropriate data collection protocols as wells as requirements

described in the appropriate assessment methods established in Assessment

Methodology for Streams and Rivers (Assessment Book, Shull and Whiteash 2021).

The protected water uses include Aquatic Life, Water Supply, Recreation and Fish

Consumption, and Special Protection.

Aquatic Life Use Monitoring

Aquatic Life Uses (ALU) include Cold Water Fishes (CWF), Warm Water Fishes (WWF),

Trout Stocking (TSF), and Migratory Fishes (MF). See 25 Pa. Code § 93.3 for

definitions and standards.

There are multiple approaches that can be considered in developing a sampling

collection plan with the objective of assessing an ALU. The first includes employing a

biological data collection protocol (Chapter 3) with a method in the Assessment Book

(Shull and Whiteash 2021). DEP currently has four benthic macroinvertebrate data

collection protocols with assessment methods. Each data collection protocol describes

criteria that will need to be evaluated to determine the appropriate protocol required for

a specific waterbody, waterbody reach, or sampling location. Each assessment method

establishes impairment thresholds and/or decision-making matrices to determine

impairment. Progressively, alternative approaches include employing a biological data

collection protocol or protocols with additional physical, chemical and/or other biological

data collections. These additional data collections, often determined during the

reconnaissance process, can be used to further assess a waterbody and used to

identify potential sources and causes of impairment.

Physical habitat data collection will be completed when employing all biological data

collection protocols. Physical habitat data can be used in conjunction with biological

assessment data or can be used independently to make ALU assessments. Guidance

on physical habitat data collection can be found in Chapter 5 of this book, or in the

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specific biological data collection protocols in Chapter 3. Specific ALU physical

assessment methodology can be found in the Assessment Book, Chapter 4.1, Physical

Habitat Assessment Method (Walters 2017). Collectors should be familiar with physical

habitat assessments and should consult associated habitat data sheets during field

reconnaissance to aid in properly identifying and locating sampling locations.

Chemical data can also be collected to characterize water quality during biological data

collection or the field reconnaissance process. Chemical data collection protocols can

be found in Chapter 4 and are required for many of the data collection protocols found

in this book. In-situ field meter data, including cross-section surveys, are highly

recommended as a screening process to determine changing water quality that may

drive differences in aquatic life. For surface waters with perceived or documented water

quality complexities (incomplete mixing, dynamic seasonal/temporal changes, etc.), and

especially on larger waterbodies (> 1000 mi2), multiple field meter data collections,

cross-section surveys, and/or discrete water chemistry data collections may be

completed multiple times targeting changing conditions throughout a period leading up

to biological data collection. A “period” is delineated by seasonality or index periods

described in the biological collection protocols and assessment methods. For example,

instream macroinvertebrate communities significantly change throughout late-May and

into June and again typically in October, depending on climatic conditions.

Consequently, collectors will usually employ macroinvertebrate collection in November

or in April. If collection is scheduled for April, multiple chemical data collection efforts

that target changing conditions beginning November and continuing through April would

provide data as to the source and/or cause of differences in biological sample results.

This information could be collected after biological data collection and after realization of

aquatic life use impairment. However, subsequent data collections may not be directly

representative of water quality conditions that caused the impairment, especially with

highly variable surface waters.

DEP also employs fish data collection, mussel data collection and periphyton (algal)

data collection protocols that can be found in Chapter 3. The latest edition of the

Assessment Book (Shull and Whiteash 2021) does include the Stream Fish

Assemblage Assessment Method (Wertz 2021d), which is designed to make

assessment determinations across Pennsylvania’s lotic surface waters. While

assessment method development is actively being pursued for fish data collection in

lentic surface waters, mussel data collection and periphyton (algal) data collection,

currently there are no assessment methods for these biological data collection

protocols, however they are employed to assess the narrative criteria and measure

changes in water quality.

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Water Supply Use Monitoring

Water Supply uses include Potable Water Supply (PWS), Industrial Water Supply

(IWS), Livestock Water Supply (LWS), Wildlife Water Supply (AWS), and Irrigation

(IRS). See 25 Pa. Code § 93.3 for definitions and standards.

Water supply use employs chemical data collection protocols including the In-Situ Field

Meter and Transect Data Collection Protocol (Hoger 2020) and the Discrete Water

Chemistry Data Collection Protocol (Shull 2017), found in Chapter 4. While 25 Pa. Code

§ 93.4 and § 96.3(c) identifies PWS as applying to all surfaces waters, § 96.3(d) states,

“(d) As an exception to subsection (c), the water quality criteria for total dissolved

solids, nitrite-nitrate nitrogen, phenolics, chloride, sulfate and fluoride established

for the protection of potable water supply [PWS] shall be met at least 99% of the

time at the point of all existing or planned surface potable water supply

withdrawals unless otherwise specified in this title”

Therefore, reconnaissance will need to identify surface potable water supply

withdrawals and sampling plans will need to target these locations, especially when

chemistry data collected involves the parameters listed in § 96.3(d).

Recreational Use and Fish Consumption Monitoring

Recreation and Fish Consumption uses include Boating (B), Fishing (F), Water Contact

(WC), and Esthetics (E). Primary objectives for recreational use and fish consumption

are Fishing (F) and Water Contact (WC). See 25 Pa. Code § 93.3 for definitions and

standards.

A sample collection plan with the objective of assessing the water contact (WC) Use will

employ Chapter 3.11, Bacteriological Data Collection Protocol (Miller 2021) and Chapter

2.5 of the Assessment Book (Shull and Whiteash 2021), Bacteriological Assessment

Method for Water Contact Sports (Miller and Whiteash 2021). Chemical data collection

protocols found in Chapter 4 of this book, including the In-situ Field Meter and Transect

Data Collection Protocol (Hoger 2020) and the Discrete Water Chemistry Data

Collection Protocol (Shull 2017), should also be considered as part of the field

reconnaissance effort.

Collecting fish consumption/fish tissue samples is typically accomplished by employing

the Fish Data Collection Protocol (Wertz 2021a) as well as the Fish Tissue Data

Collection Protocol (Wertz 2021c), Chapter 3, and samples are often collected as a

secondary objective during community surveys. Other acceptable collection procedures

include seine, gill net, rotenone, angling, etc. Refer to Chapter 2 of the Assessment

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Book (Shull and Whiteash 2021), Fish Tissue Consumption Assessment Method (Wertz

2021b) for additional requirements that will need to be addressed in the sample

collection plan.

Special Protection Monitoring

Special Protection uses include High Quality Waters (HQ) and Exceptional Value

Waters (EV). See 25 Pa. Code § 93.3 for definitions and standards.

Developing a sample collection plan for special protection waters will include employing

a biological data collection protocol (Chapter 3) with an assessment method. Each data

collection protocol describes criteria that will need to be evaluated to determine the

appropriate protocol required for a specific waterbody, waterbody reach, or sampling

location. When appropriate, biological assessment methods will include, index periods

and impairment thresholds and decision matrices specific to special protection waters

that will need to be addressed in the sample collection plan. Otherwise, sample

collection plan development is consistent with aquatic life use monitoring.

Stream Redesignation Evaluation Monitoring

Stream redesignation evaluations are initiated to determine the existing and designated

uses of a given waterbody. The terms “existing use” and “designated use” are defined at

25 Pa. Code § 93.1. Evaluations can be initiated as part of a petition process, permitting

process, or due to the results of a use assessment reconnaissance or survey that

indicates an existing use may be different than the designated use. Redesignation

evaluations can lead to redesignation to a more restrictive use, a less restrictive use, no

change to the use, or a demonstration of “natural quality” as defined at 25 Pa. Code §

93.1.

A sample collection plan for redesignation evaluations will include and refer to

information at 25 Pa. Code § 93.4b, ‘Qualifying as High Quality or Exceptional Value

Waters’ and the DEP Water Quality Antidegradation Implementation Guidance (DEP

2003). This includes the application of biological and chemical data collection protocols.

Physical habitat assessments will also be completed when employing biological data

collection protocols. Thermal components of ALU (WWF, TSF, and CWF) are evaluated

using fish community data and will employ the Fish Data Collection Protocol (Wertz

2021c) found in Chapter 3.

Point of First Surface Water Use Monitoring

Point of first use surveys are completed as part of the evaluation and permitting of

wastewater discharges to intermittent and ephemeral streams, drainage channels and

swales, and storm sewer. The point of first surface water use establishes where

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Chapter 93 Water Quality Standards (WQS) must be attained. Collectors will need to

become familiar with DEP technical guidance document 391-2000-014, Policy and

Procedures for Evaluating Wastewater Discharges to Intermittent and Ephemeral

Streams, Drainage Channels and Swales, and Storm Sewers (DEP 2000). Point of first

use surveys will employ biological data collection protocols found in Chapter 3.

Cause and Effect Monitoring

A cause and effect sampling design is employed to investigate possible relationships

between point or nonpoint sources of conventional pollutants and known or suspected

instream water quality problems through the collection and analysis of biological,

physical, and chemical data. These surveys are designed primarily to monitor the

effectiveness of permitted treatment facilities but are also used to investigate reported

or suspected water quality impacts for nonpoint source and other pollution sources.

Cause and Effect Surveys (Lookenbill and Wertz 2021) found in Chapter 2.1 includes

specific survey design requirements. Cause and effect monitoring will employ biological,

chemical, and physical data collection protocols found in this book.

Water Quality Network Monitoring

The Pennsylvania Water Quality Network (WQN) is a statewide, fixed station water

quality sampling system operated by DEP. It is designed to assess both the quality of

Pennsylvania’s surface waters and the effectiveness of the water quality management

program by accomplishing four basic objectives:

• Monitor temporal water quality trends in major surface streams throughout

Pennsylvania.

• Monitor temporal water quality trends in selected reference waters.

• Monitor the trends of nutrient and sediment loads in the major tributaries entering

the Chesapeake Bay.

• Monitor temporal water quality trends in selected Pennsylvania lakes.

Data collection is a collaborative effort between the US Geological Survey (USGS) PA

Water Science Center, Susquehanna River Basin Commission (SRBC), and DEP. WQN

monitoring employs routine biological data collection (primarily benthic

macroinvertebrate data collection, chemical/physical data collection, and stream

discharge data collection. Appendix D contains Pennsylvania’s Surface Water Quality

Monitoring Network (WQN) Manual (Pulket and Lookenbill 2020) which details the

network’s sampling design, station locations, and the water chemistry analytes

collected.

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Protocol, Method, and Standards Development

The DEP Water Quality Division (WQD) is responsible for developing surface water

data collection protocols, water quality criteria and protected water uses, assessment

methods, Pennsylvania’s biannual integrated monitoring and assessment report, and

Total Maximum Daily Loads (TMDLs) that allow impaired waters to meet WQS. Data

collection to meet an objective of protocol, method or WQS development is unique from

other data collection objectives. The ultimate purpose is to implement The Pennsylvania

Clean Streams Law and the Federal Clean Water Act. To achieve this, at a statewide

level, WQS are developed to establish the criteria and protected uses that surface

waters must meet. Data collection protocols and assessment methods are developed,

at a statewide level, to collect consistent data that will be used to determine if surface

waters meet WQS. In addition, data collection protocols also need to be applicable to

other data collection objectives.

Sampling design for a development objective will pursue the range of possible water

quality conditions across the State. Reconnaissance will focus on a range of conditions

and attributes that describe this range and not necessarily the potential to delineate a

water quality condition or the source or cause on a waterbody. A range of conditions will

be qualified by identifying classification of waterbodies. Attributes that are selected to

describe a range of water quality conditions will rely on published literature.

Development objectives, particularly the development of standards, may also rely on

laboratory investigations.

LITERATURE CITED

Brickner, M. (editor). 2020. Macroinvertebrate laboratory subsampling and identification.

Chapter 3.6, pages 32–42 in M. J. Lookenbill and R. Whiteash (editors). Water

quality monitoring protocols for streams and rivers. Pennsylvania Department of

Environmental Protection, Harrisburg, Pennsylvania.

DEP. 2000. Policy and procedures for evaluating wastewater discharges to intermittent

and ephemeral streams, drainage channels and swales, and storm sewers.

Pennsylvania Department of Environmental Protection, Harrisburg, Pennsylvania

(Available from:

http://www.depgreenport.state.pa.us/elibrary/GetDocument?docId=7790&DocNa

me=391-2000-014.pdf)

DEP. 2003. Water quality antidegradation implementation guidance. Pennsylvania

Department of Environmental Protection, Harrisburg, Pennsylvania. (Available

from: http://www.depgreenport.state.pa.us/elibrary/GetFolder?FolderID=4664)

Hoger, M. 2020. In-situ field meter and transect data collection protocol. Chapter 4.1,

pages 2–8 in M. J. Lookenbill and R. Whiteash (editors). Water quality monitoring

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protocols for streams and rivers. Pennsylvania Department of Environmental

Protection, Harrisburg, Pennsylvania.

Lookenbill, M. J. 2020a. Combined sewer overflow (CSO) compliance monitoring.

Chapter 2.1, pages 14–23 in M. J. Lookenbill and R. Whiteash (editors). Water

quality monitoring protocols for streams and rivers. Pennsylvania Department of


Lookenbill, M. J. 2020. Sampling design and planning. Chapter 2, pages 1–25 in M. J.

Lookenbill and R. Whiteash (editors). Water quality monitoring protocols for

streams and rivers. Pennsylvania Department of Environmental Protection,

Harrisburg, Pennsylvania.

Lookenbill, M. J. and Wertz, T. A. (editors). 2021. Cause and effect surveys. Chapter

2.1, pages 7–15 in M. J. Lookenbill and R. Whiteash (editors). Water quality

monitoring protocols for streams and rivers. Pennsylvania Department of


Miller, S. (editor). 2021. Bacteriological data collection protocol. Chapter 3.11, pages

142–158 in M. J. Lookenbill and R. Whiteash (editors). Water quality monitoring



Miller, S., and R. Whiteash. 2021 (editors). Bacteriological assessment method for

water contact sports. Chapter 2.5, pages 75–79 in D. R. Shull, and R. Whiteash

(editors). Assessment methodology for streams and rivers. Pennsylvania

Department of Environmental Protection, Harrisburg, Pennsylvania.

Pulket, M., and M. J. Lookenbill (editors). 2020. Pennsylvania’s surface water quality

monitoring network manual. Appendix D, pages 1–71 in M. J. Lookenbill, and R.

Whiteash (editors). Water quality monitoring protocols for streams and rivers.

Pennsylvania Department of Environmental Protection, Harrisburg,

Pennsylvania.

Shull, D. R. 2017. Discrete water chemistry data collection protocol. Chapter 4.2, pages

9–24 in M. J. Lookenbill, and R. Whiteash (editors). Water quality monitoring



Shull, D. R., and R. Whiteash (editors). 2021. Assessment methodology for streams and

rivers. Pennsylvania Department of Environmental Protection, Harrisburg,

Pennsylvania.

Wertz, T. A. (editor). 2021a. Fish data collection protocol. Chapter 3.7, pages 43–62 in

M. J. Lookenbill, and R. Whiteash (editors). Water quality monitoring protocols for



Wertz, T. A. (editor). 2021b. Fish tissue consumption assessment method. Chapter 2.6,

pages 80–87 in D. R. Shull, and R. Whiteash (editors). Assessment methodology

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for streams and rivers. Pennsylvania Department of Environmental Protection,


Wertz, T. A. (editor). 2021c. Fish tissue data collection protocol. Chapter 3.8, pages 63–

73 in M. J. Lookenbill, and R. Whiteash (editors). Water quality monitoring



Wertz, T. A. 2021d. Stream fish assemblage assessment method. Chapter 2.7, pages

88–107 in D. R. Shull, and R. Whiteash (editors). Assessment methodology for



Walters, G. 2017. Physical habitat assessment method. Chapter 4.1, pages 2–6 in D. R.

Shull, and R. Whiteash (editors). Assessment methodology for streams and


Pennsylvania.

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Prepared by:

Josh Lookenbill






2018

Edited by:

Josh Lookenbill






2020

2-3

TYPES OF SAMPLING DESIGN

A sampling design must be chosen in order to form a data set that is appropriate to

describe the desired objective. The two general sample design categories are

probabilistic (also called “statistical” or “random”) sampling designs and judgmental or

targeted sampling designs (USEPA 2002a).

Probabilistic Sampling

A probabilistic or probability-based sampling design utilizes randomization in the

selection of sample locations (USEPA 2002a). A probabilistic sampling design requires

accurate information about the population being sampled to ensure that a consistent

distribution of the population is sampled. The sampling location selection process must

consider population characteristics (Strahler stream order, landuse, number and

location of point source discharges, etc.) and must be weighted accordingly. There are

various ways that sites can be picked for a probabilistic monitoring study. One approach

is a Generalized Random Tessellation Stratified (GRTS) design procedure. This

procedure was employed for DEP’s recreational use monitoring and statewide statistical

survey requirements. GRTS provides a random sample that is balanced spatially. The

sites are chosen using an R-program package called ‘spsurvey’ (Kincaid and Olsen

2011). The package ‘spsurvey’ also helps analyze the results and can give estimations

of the proportion of samples in each category analyzed, along with standard error and

total estimates (Kincaid 2012a, Kincaid 2012b). Other procedures may be employed to

choose sites statistically; however, without a detailed characterization and classification,

a probabilistic sampling design does not necessarily provide for detailed assessment

delineations and is not the preferred sampling design.

Targeted Sampling

DEP’s primary monitoring objective is to collect data to assess protected water uses of

specific water segments or reaches, where most other states assess larger watershed

units. DEP believes the targeted “judgment-based” sampling design is the most suited

procedure to assess WQS including uses of specific water segments or reaches. A

targeted or judgmental sampling design provides for the selection of sample locations

based on professional judgment and does not utilize randomization. This approach

requires intense reconnaissance and information gathering but can result in an

accurately delineated assessment and is the preferred sampling approach for DEP

monitoring protocols and subsequent assessment methods.

Reconnaissance includes the delineation of the targeted waterbody or basin and the

gathering of historical and other information. A review of any gathered data or

information may influence the geographical extent of the survey. Aerial photography,

2-4

topographic maps, and/or geographical information system (GIS) information are used

to delineate the targeted waterbody, identify major tributaries, characterize land use and

any potential sources of nonpoint source impacts (agriculture, urban areas, etc.),

potential sources of point source impacts (industrial discharges, sewage treatment

discharges, combined sewer overflow (CSO discharges, etc.), geology, and soil

characteristics. Historical information includes past water quality data and associated

use assessments. Other information can include designated and existing water uses

and ongoing water quality monitoring like the DEP Water Quality Network (WQN) and

US Geological Survey (USGS) stream gaging network.

Sampling sites and locations are positioned to account for changes in water quality due

to influences such as major tributaries, point and nonpoint source impacts, land use

changes, soil characteristics, and geology. Additional samples are collected at the limits

of these changes to effectively “bracket” potential sources of water quality differences.

Initial sampling site placement should target 3rd order or higher waterbodies. Sampling

sites located on 1st and 2nd order waterbodies are chosen based on the variability of

potential changes to water quality, but not all 1st and 2nd order waterbodies will have

sampling sites located on them. It is important to identify 1st and 2nd order tributaries that

may be ephemeral or intermittent, as assessment methods are typically not developed

to account for this variable. Also, the minimum length of any use assessment unit is ½

mile. Any impact delineated in length that is less than ½ mile is considered a localized

impact, potentially a compliance issue, but not a non-attainment of use. Additional field

reconnaissance is highly recommended to verify potential sources of water quality

impacts. Field reconnaissance will include visual observations and physical habitat

evaluations (see Chapter 5) and can also include chemical data collection and/or field

meter and cross-section survey data collection (see Chapter 4). Cross-section surveys

performed using a clean and calibrated field meter that collects water temperature,

specific conductance, dissolved oxygen, pH, and – preferably – turbidity are required to

determine if major water quality influences exist at the sampling location.

SAMPLING PLANS

A sample collection plan is recommended before commencing a sampling project. This

should include the primary and any secondary objectives, subsequent monitoring

protocols and assessment methods that will be employed to meet objectives, tentative

sampling/site location(s), timing and frequency of samples, media of interest (water,

sediment, soil, macroinvertebrates, etc.), parameters to sample, QA and Quality Control

(QC) plans, sampling supplies and equipment, and any other notes or comments

pertaining to the project. More information on developing Project Plans can be found in

2-5

United States Environmental Protection Agency (USEPA) resources (USEPA 2002a,

USEPA 2002b, and USEPA 2006).

A sample collection plan should describe how the data will be used. Analyses and data

interpretation should be anticipated in advance. A collector should consider if an

objective is to defend an enforcement action or recommend regulatory changes. Also,

consider what criteria may be used during its interpretation. DEP has numeric and

narrative water quality criteria documented in 25 Pa. Code Chapter 93, Water Quality

Standards. Numeric criteria allow for screenings during sampling. In some cases, there

may be no numeric criteria for a substance or sampling matrix. For example, DEP has

no numeric sediment criteria at this time. However, in order to apply these or other

results that do not have numeric criteria, one can refer to the 25 Pa. Code § 93.6:

“(a) Water may not contain substances attributable to point or nonpoint source

discharges in concentration or amounts sufficient to be inimical or harmful to the

water uses to be protected or to human, animal, plant or aquatic life. and (b) In

addition to other substances listed within or addressed by this Chapter, specific

substances to be controlled include, but are not limited to, floating materials, oil,

grease, scum and substances that produce color, tastes, odors, turbidity or settle

to form deposits.”

The numeric criteria, narrative criteria, or biological standard to be measured should be

described prior to study commencement.

LITERATURE CITED

Kincaid, T. M., and A. R. Olsen. 2011. spsurvey: Spatial Survey Design and Analysis. R

package version 2.2.

Kincaid, T. M. 2012a. Analysis of a GRTS survey design for a linear resource. CRAN

– R project.

Kincaid, T. M. 2012b. GRTS survey designs for a linear resource. CRAN – R project.

USEPA. 2002a. Guidance on choosing a sampling design for environmental data

collection for use in developing a quality assurance project plan.

EPA/240/R02/005. EPA QA/G-5S. U.S. Environmental Protection Agency, Office

of Environmental Information, Washington, DC.

USEPA. 2002b. Guidance for quality assurance project plans. EPA/240/R-02/009. EPA

QA/G-5. U.S. Environmental Protection Agency, Office of Environmental

Information, Washington, DC.

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USEPA. 2006. Guidance on systematic planning using the data quality objectives

process. EPA/240/B-06/001. EPA QA/G-4. U.S. Environmental Protection

Agency, Office of Environmental Information, Washington, DC.

2-7

2.1 CAUSE AND EFFECT SURVEYS

2-8

Prepared by:

Josh Lookenbill






2018

Edited by:

Josh Lookenbill and Timothy Wertz






2021

2-9

INTRODUCTION

Cause and effect surveys are designed to investigate possible relationships between

point or nonpoint sources of conventional pollutants and known or suspected instream

water quality problems through the collection and analysis of biological, physical, and

chemical data. This protocol was developed to establish and standardize cause and

effect survey procedures and provide guidance to DEP staff for conducting such

surveys. The protocol should be used in conjunction with other DEP data collection

protocols and assessment methods.

These surveys are performed primarily to monitor the effectiveness of permitted

treatment facilities but are also used to investigate reported or suspected water quality

impacts from nonpoint source and other pollution sources. Since such discharges exist

on a wide variety of stream and river habitats, the sampling design and type of sampling

gear used are dependent on stream-type, site-specific conditions, and the nature of the

discharge under investigation. DEP staff are responsible for sampling design and any

modifications to sampling design due to unforeseen site-specific conditions.

SAMPLING DESIGN

The sampling design for a cause and effect survey requires a minimum of two sampling

stations. One station is placed upstream from the subject discharge(s) or impact(s) to

serve as a control or reference condition and at least one station is placed in a

potentially impacted zone downstream. Additional sampling stations may be necessary,

either placed downstream to define zones of impact and recovery; upstream to bracket

multiple discharges; or across a stream transect in wider waterbodies. For point

sources, observations in the immediate vicinity of a discharge may also be useful.

Observations in the immediate vicinity of a discharge should include:

• Floating solids, scum, sheen, or substances that result in observed deposits in

the receiving water (a small amount of foam that rapidly dissipates is typical);

• Oil and grease in amounts that cause a film or sheen upon or discoloration of the

surface waters or adjoining shoreline, or that exceed 15 mg/l as a daily average

or 30 mg/l at any time (or lesser amounts if specified in the respective permit);

• Substances in concentration or amounts sufficient to be inimical or harmful to

the water uses to be protected or to human, animal, plant, or aquatic life;

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• Foam or substances that produce an observed change in the color, taste, odor,

or turbidity of the receiving water, unless those conditions are otherwise

controlled through effluent limitations or other requirements in the respective

permit.

Any noted observations may or may not indicate non-attainment of WQS or a violation

of associated permit conditions. Regulatory requirements and permit conditions

(including 25 Pa. Code §§ 93.6, 92a.41(c), 95.2(2)(i)) should be consulted.

With the exception of observations previously described, stations sampled downstream

of the discharge(s) or nonpoint sources should not be placed in the immediate vicinity of

the discharge or nonpoint source, but instead should generally be located a distance

downstream in a potentially impacted zone. Selecting downstream station locations is

unrelated to “criteria compliance times” used to generate National Pollutant Discharge

Elimination System (NPDES) permit limits. Similarly, it is not necessary to restrict the

location of downstream stations to points at or downstream of the point of complete mix,

as defined as the point where water quality and/or other characteristics are

homogenous across a transect. If a tributary or other compromising influence is located

just downstream of the discharge or source of potential impact, which would not make

an upstream/downstream comparison directly applicable, DEP staff may need to modify

the sampling design to account for differences in water quality that are not necessarily

due to the discharge or source of potential impact.

Complete mix occurs rapidly in smaller streams at most flow conditions and it is typically

appropriate to locate downstream stations in smaller streams at a point of complete mix.

In larger streams or rivers, a plume of effluent could extend a significant distance

laterally and longitudinally. An indicator station may be located at or downstream of this

point recognizing that any chemical, physical, or biological sampling could be conducted

as a composite across the transect, but effects within the plume will also be assessed if

a significant portion of the receiving water is impacted. The number and placement of

any in-situ collections across a transect and subsequent compositing should be

completed according to Chapter 4.1, In-situ Field Meter and Transect Data Collection

Protocol (Hoger 2020), or equal-width-increment or equal-discharge-increment

procedures (USGS 2006). When collecting biological samples, the equal-width-

increment procedure can be considered (i.e., effort physically distributed equally across

the/a transect), but the sample should be collected in a manner such that the sample

represents the waterbody across the transect and targets best available habitat

according to the DEP protocols and methodologies.

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If the downstream station is placed where homogenous conditions do not exist,

additional downstream cross-section surveys and sampling stations may be necessary

to characterize the effect. Any critical habitat of threatened or endangered species, as

defined by the United States Fish and Wildlife Service, or any rare or endemic

ecological community types, as defined by the Pennsylvania Natural Heritage Program,

or any migration impediments must be identified within the defined plume or within the

vicinity of the plume. The water quality and its effect on identified habitat or ecological

communities and/or WQS will be assessed specific to identified habitat or ecological

communities.

Control or reference sampling stations may be placed at any point upstream from the

discharge or source where there is no potential impact from the discharge at any river

flow condition. In a low-gradient situation such as a pool, it may be most appropriate to

locate the reference station far enough upstream from the discharge to preclude

possible effects from pooling effluent during low river flow conditions. If there is an

intake structure present, it may be most appropriate to locate the reference station

upstream from the intake structure to preclude possible effects from recirculating

effluent during low river flow conditions.

In some instances, if upstream conditions do not adequately represent control or

reference conditions, it may be necessary to sample a separate waterbody for reference

purposes. Ideally, this reference station would be selected within the same watershed or

if no available reference station is found, an appropriate station should be in an adjacent

watershed. Water quality impacts to impounded waters present at least the

compounding effect the impoundment itself has on water quality and water quality

indicators and may prevent traditional upstream/downstream sampling designs from

detecting changes in water quality due to impacts other than the impoundment. To

reduce or eliminate compounding sources of variability, physical habitat conditions

should be as similar as possible, for each segment or separate waterbody selected for

sampling. The following data will be collected at all sampling stations: benthic

macroinvertebrates, habitat assessment, field measurements or water chemistry,

channel cross-section and stream flow (as needed), cross-section surveys (as needed),

bacteria (as needed), and fish (as needed).

DATA COLLECTION

Benthic Macroinvertebrates (required)

For wadeable freestone streams, limestone-influenced streams, low gradient streams,

or large semi-wadable rivers, benthic macroinvertebrate samples are collected utilizing

protocols detailed in Chapter 3 of this book. Each benthic macroinvertebrate data

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collection protocol describes what will need to be evaluated to determine which protocol

to use. Data collection protocols include semi-quantitative (population densities derived

by estimation), qualitative (population densities not calculated), and quantitative

(population densities derived by measurement) protocols. The survey data needs will

determine which protocol is most suitable. In most instances, semi-quantitative data

collection protocols are preferred and will meet most data requirements. Other optional

data collection protocols include qualitative and quantitative. Qualitative protocols

(Chapter 3) are appropriate Cause and Effect Survey protocols. Qualitative protocols

may be more appropriate than quantitative or semi-quantitative protocols, and may be

used in place of, or in conjunction with semi-quantitative data collection when the survey

requires a more immediate result or the targeted surface water is not characteristic of

surface waters used to develop a method or index. Qualitative surveys are also

appropriate as preliminary evaluations immediately following acute pollution events

where the impact to the biological community doesn’t occur for a period of one to three

weeks. Qualitative surveys could be used to determine the timing of subsequent

quantitative or semi-quantitative surveys.

Regardless of the data collection protocol used, sampling stations consist of a control or

background station placed upstream from the discharge(s) and at least one affected or

impacted station downstream from the discharge(s) in the best available riffle and run

habitat. When multiple discharges are present, sampling stations are placed between

discharges to characterize the effect of each input. Physical variables of all sample

stations should be matched as closely as possible between background and impacted

stations to minimize or eliminate the effects of compounding variables. Sample points

are placed to obtain a representative benthic sample and to avoid over sampling of

clustered populations.

DEP developed an index of biotic integrity (IBI) for benthic macroinvertebrate

communities collected via semi-quantitative protocols in Pennsylvania’s wadeable,

freestone, riffle-run streams. See Chapter 2 of the Assessment Book (Shull and

Whiteash 2021). Through direct quantification of biological attributes along a gradient of

ecosystem conditions, this IBI measures the extent to which anthropogenic activities

compromise a stream’s ability to support healthy aquatic communities (Davis and Simon

1995) and can be used to compare control or reference conditions versus impacted

conditions. DEP’s latest IBIs describe precision estimates for temporal variability

(...whether a site’s biological condition has improved or degraded over time) and

intersite variability (…whether a site’s biological condition is improved or degraded when

compared to a nearby site). Samples collected for cause and effect surveys should be

collected on the same day to eliminate the need to consider temporal variability.

Intrasite variability, as determined by the IBI for benthic macroinvertebrates applies to

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samples collected from a single site/station (within 100 meters). Cause and effect

survey upstream and downstream stations are collected from separate sites and

therefore an intersite precision estimate was developed and should be applied to

compare upstream, downstream and recovery zone sites/stations. Cause and effect

upstream or control versus downstream or recovery stations collected using the latest

semi-quantitative protocols for wadeable, freestone, riffle-run streams with IBI scores

greater than the intersite precision estimate will be considered impacted. The

appropriate intersite precision estimate is determined by DEP Water Quality Division,

Assessment Section. Follow-up surveys may be conducted when small IBI score

differences are found between control and impact sites, to confirm, or re-evaluate initial

cause and effect survey results.

Fish (optional)

For most cause and effect surveys semi-quantitative fish sampling should be

considered. See Chapter 3.7, Fish Data Collection Protocol (Wertz 2021a). The

objective is to acquire a representative sample of the fish population by sampling all

physical stream habitats in relative proportion to their availability.

DEP developed Stream Fish Assemblage Assessment Method (Wertz 2021b) using a

Thermal Fish Index (TFI) for fish communities collected via semi-quantitative protocols

in Pennsylvania’s lotic streams and rivers. The assessment method provides additional

guidance for initiating and evaluating cause and effect surveys.

If fish kills are a component of a cause and effect survey investigation, quantitative

procedures would be required in cases where economic damages may need to be

calculated resulting from incidents causing fish kills. In these instances, to support

fish/aquatic life penalties, the required fish/aquatic life data should be collected in a

manner consistent with The American Fisheries Society (Southwick and Loftus 2017) or

Pennsylvania Fish and Boat Commission (PFBC) fish kill survey procedures. In many

cases, it may be more practical to coordinate field sampling activities with the regional

PFBC Fisheries Management staff.

Bacteria (optional)

Because of the survey and cost complexities imposed by bacterial sampling (sample

frequency and short holding times), bacteria sample collection is an optional

consideration limited to when sanitary impacts from discharges are suspected. Samples

for bacteriological analysis are collected to define the sanitary significance of point and

nonpoint sources and assess the use attainment status of stream segments for potable

water supply and recreational uses. The samples are collected using protocols outlined

in Chapter 3 of this book.

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Habitat Evaluation (required)

A habitat evaluation is conducted on a measured 100-meter reach of stream at a

minimum for wadeable surface waters according to Chapter 5.1, Stream Habitat Data

Collection Protocol (Lookenbill 2017) in this Monitoring Book and Chapter 4.1, Physical

Habitat Assessment Method (Walters 2017) in the Assessment Book. The habitat

evaluation process involves rating twelve parameters for riffle/run prevalent waterbodies

or nine parameters for low gradient waterbodies as optimal, suboptimal, marginal, or

poor by using a numeric value (ranging from 20-0).

Field Measurements and Water Chemistry (required)

Detailed field observations on land use and potential sources of pollution are recorded

on field data collection forms or in field books. Dissolved oxygen, pH, specific

conductance, and temperature are measured in the field using hand-held meters

calibrated according to manufacturer specifications and the latest DEP protocols found

in Chapter 4 of this book.

One-time grab samples are collected, at a minimum, from a control or background

station upstream from the discharge, from the discharge, and from at least one

downstream affected or impacted station when evaluating point source discharges.

Point source discharges that do not continuously discharge or those that discharge

subsurface may prevent grab samples from being collected. In these situations,

Discharge Monitoring Reports (DMRs) that document the chemical violation of the

NPDES permit would suffice. For nonpoint discharges, grab samples are collected

upstream of the impacted segment and from within the impacted segment.

Discharge (optional)

Discharge data is collected as needed according to the latest version of USGS

Techniques and Methods book 3, chap. A8, Discharge Measurements at Gaging

Stations (Turnipseed and Sauer 2010).

LITERATURE CITED

Davis, W. S. and T. P. Simon. 1995. Introduction to biological assessment and criteria:

Tools for water resource planning and decision making. Pages 3–6 in W. S.

Davis, and T. P. Simon (editors). CRC Press, Boca Raton.

Hoger, M. 2020. In-situ field meter and transect data collection protocol. Chapter 4.1,




2-15

Lookenbill, M. J. (editor). 2017. Stream habitat data collection protocol. Chapter 5.1,






Pennsylvania.

Southwick, R. I., and A. J. Loftus. 2017. Investigations and monetary values of fish and

freshwater mussel kills. American Fisheries Society. ISBN 978-1-934874-47-9.

Turnipseed, D. P. and V. B. Sauer. 2010. Discharge measurements at gaging stations:

United States Geologic Survey. Book 3, chapter A8. in National field manual for

the collection of water quality data: Techniques and methods. U.S. Department of

the Interior, U.S. Geological Survey, Reston, VA. (Available online at:

https://pubs.usgs.gov/twri/twri3a8/.)

USGS. 2006. Collection of water samples. Book 9, chapter A4 (version 2.0). in National

field manual for the collection of water quality data: Techniques and methods.

U.S. Department of the Interior, U.S. Geological Survey, Reston, VA. (Available

online at: https://www.usgs.gov/mission-areas/water-resources/science/national-

field-manual-collection-water-quality-data-nfm?qt-science center objects=0#qt-

science center objects.)




Pennsylvania.

Wertz, T. A. 2021a. Fish data collection protocol. Chapter 3.7, pages 43–62 in M. J.

Lookenbill, and R. Whiteash (editors). Water quality monitoring protocols for



Wertz, T. A. 2021b. Stream fish assemblage assessment method. Chapter 2.7, pages

88–107 in D. R. Shull, and R. Whiteash (editors). Assessment methodology for



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2.2 COMBINED SEWER OVERFLOW (CSO) COMPLIANCE DATA COLLECTION

2-17

Prepared by:

Josh Lookenbill






2020

2-18

INTRODUCTION

CSOs are unique in the way they are regulated and so require unique sampling plans to

meet the demands of compliance monitoring. The Federal Register details the

requirements of CSO policy and monitoring requirements. Specifically, 59 FR 18688

explains:

“The CSO policy represents a comprehensive national strategy to ensure

that municipalities, permitting authorities, water quality standards

authorities and the public engage in a comprehensive and coordinated

planning effort to achieve cost effective CSO controls that ultimately meet

appropriate health and environmental objectives. …State water quality

standards authorities will be involved in the long-term CSO control planning

effort as well. The water quality standards authorities will help ensure that

development of the CSO permittees’ long-term CSO control plans are

coordinated with the review and possible revision of water quality standards

on CSO-impacted waters.”

59 FR 18696, under section 2 explains:

“Once the permittee has completed development of the long-term CSO

control plan and the selection of the controls necessary to meet CWA

requirements has been coordinated with the permitting and WQS

authorities, the permitting authority should include, in an appropriate

enforceable mechanism, requirements for implementation of the long-term

CSO control plan as soon as practicable. Where the permittee has selected

controls based on the "presumption" approach described in Section II.C.4,

the permitting authority must have determined that the presumption that

such level of treatment will achieve water quality standards is reasonable in

light of the data and analysis conducted under this Policy.”

Additionally, the Phase II permit should contain:

“A requirement to implement, with an established schedule, the approved

post-construction water quality assessment program including

requirements to monitor and collect sufficient information to demonstrate

compliance with WQS and protection of designated uses as well as to

determine the effectiveness of CSO controls.”

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USEPA’s draft CSO Post Construction Compliance Monitoring Guidance (USEPA 2011)

also recommends the information to be collected for monitoring CSO compliance,

consistent with the Federal Register. This guidance states that CSO Monitoring

programs should include both CSO effluent and ambient in-stream monitoring. USEPA’s

draft guidance also suggests, where appropriate, biological assessments, toxicity

testing, and sediment sampling are also implemented.

Based on the information found in the Federal Register and USEPA’s guidance (USEPA

2011), designing a sampling plan for CSO compliance monitoring involves answering

two important questions:

1. Does the receiving waterbody meet attainment thresholds?

2. Is the CSO causing or contributing to a WQS impairment?

These two questions are not the same, because the sampling plan will be different for

each question. To answer the first question, sampling must be designed in accordance

with making protected use assessment determinations, which are determinations on the

entire waterbody, or on defined zones as discussed in the Assessment Book (Shull and

Whiteash 2021). The assessment determination process inherently avoids collecting

data at or very near single discharge points, because of the possibility of using results

from one discharge to make an assessment determination on the entire waterbody.

Conversely, the second question requires that the sampling plan incorporates sampling

at or very near single points of discharge to determine contribution to a potential WQS

impairment. It is important to note that sampling must be conducted to answer both

questions regardless of an assessment determination that shows attainment of WQS

upstream and downstream of the CSO.

SAMPLING DESIGN

Sampling design for CSO compliance requires a combination of use assessment and

cause and effect sampling design. Both sampling designs typically include biological,

physical, and chemical data collection. To evaluate any impacts from CSOs, biological

and chemical evaluations will be the primary targets. Biological data collection will

specifically require bacteriological monitoring. Chemical data collections will require at

least discrete water chemistry monitoring. In addition, performance monitoring of the

combined sewer system(s) (CSS) will also be conducted.

All monitoring plans will be submitted to DEP for approval prior to being implemented. A

final report will be submitted within 90 days following the collection of the last samples

for DEP review.

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DATA COLLECTION

Does the receiving waterbody meet attainment thresholds?

Bacteria (required)

Bacteria data collection is conducted according to protocols outlined in Chapter 3 of this

book during the swimming season (May 1st through September 30th). Assessment

determinations will be conducted on upstream and downstream segments of the CSO

or multiple CSOs if there is not the opportunity to sample between according to methods

outlined in Chapter 2 of the Assessment Book (Shull and Whiteash 2021).


Data collection for other parameters identified in the CSO Control Policy are conducted

according to the latest DEP protocols found in Chapter 4 of this book. Assessment

determinations will be conducted on upstream and downstream segments of the CSO

or multiple CSOs if there is not the opportunity to sample between according to methods

outlined in Chapter 3 of the Assessment Book (Shull and Whiteash 2021).

Is the CSO causing or contributing to a WQS impairment?

Bacteria (required)

Bacteria data collection is conducted according to procedures outlined in Chapter 3 of

this book for at least 1 year (4 seasons) as follows:

• During each month of the recreational season (May 1st through September 30th),

samples are collected upstream and downstream of each outfall (or groups of

outfalls as appropriate).

o Samples must be analyzed for fecal coliform1 and E. coli.2

o 5 samples must be collected under “normal” recreating conditions.

o 5 samples must be collected under other, critical conditions to capture

high flows, storm events, etc. when CSOs are discharging.

o Corresponding qPCR samples should also be collected for each set of

samples. A minimum of 1 sample per monthly set is recommended, but

additional samples are encouraged as they will better characterize the

source(s) of bacteria to determine whether the CSO is contributing to the

bacterial load.

1, 2 At the time of publication, fecal coliform limitations are referenced at 25 Pa. Code § 92a.47 and E. coli

criteria are listed at § 93.7.

2-21

• During each month of the non-recreational season (October 1st through April

30th), samples are collected at the same upstream and downstream locations

established during the recreational season following the same protocol.

o Samples must be analyzed for fecal coliform1 and E. coli.2

o 5 samples must be collected in accordance with data collection protocols

during normal/low flow conditions.

o 5 samples must be collected under other, critical conditions to capture

high flows, storm events, etc.

o Corresponding qPCR samples should also be collected for each set of

samples. A minimum of 1 sample per monthly set is recommended, but

additional samples are encouraged as they will better characterize the

source(s) of bacteria to determine whether the CSO is contributing to a

use impairment or impact to the surface water.

1, 2 At the time of publication, fecal coliform limitations are referenced at 25 Pa. Code § 92a.47 and E. coli

criteria are listed at § 93.7.


Data collection for other parameters identified in the CSO Control Policy are conducted

according to the latest DEP protocols found in Chapter 4 of this book for at least 1 year

(4 seasons) as follows:

• At minimum, these parameters include BOD, TSS, ammonia-N, specific

conductance, pH, temperature and DO as well as any other parameters

reasonably expected to be in the discharge.

• A minimum of 1 sample should be collected during each CSO discharge event

throughout the 1-year sampling period unless dangerous conditions prohibit

sample collection.

CSS Performance Monitoring (required)

The purpose of performance monitoring is to document frequency and duration of CSO

activations. A specific CSO activation event definition should be defined for each CSS.

Performance monitoring includes the characterization of CSO activations and the

collection of discharge data at strategic locations throughout and rainfall data within the

CSS catchment area(s).

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CASE STUDY

The following case study demonstrates the need to collect data using both sampling

designs. Briefly, this study demonstrates adequate sampling design to answer question

2 (Is the CSO contributing to a WQS impairment?), but there is insufficient information

to determine the answer to question 1 (Does the receiving waterbody meet attainment

thresholds?). Therefore, more data needs to be collected to make use assessment

determinations on stream segments upstream and downstream of the CSO.

A Post-Construction Monitoring Plan and subsequent results from 2014–

2017 were provided to a DEP regional office. The plan outlined routine

surface water monitoring that took place at 7 sites, upstream and

downstream of discharges. All samples were collected from the shoreline.

Sites were identified as Site 1 through Site 8, moving upstream to

downstream, respectively (Site 5 was discontinued). Site 1 was located

upstream of all urbanized area and all CSO outfalls. Sites 2 through 6

were located appropriately to measure the contribution of several CSO

outfalls and bracket one tributary influence. Site 7 was located

downstream of all CSO outfalls, an urban influenced tributary, and a

sanitary sewer outfall. Site 8 was located downstream of the entire

urbanized area.

Sampling frequency was five times per month during the swimming

season (May 1st – September 30th). The goal of this approach was to

capture both wet and dry weather events, if possible. Samples were

collected on the first four Tuesdays of each month during the swimming

season with a fifth “floating” sample taken each month. There was no data

collected during the non-swimming season, and no qPCR samples were

collected.

A review of the data revealed that 5-base flow (0 – 0.25” rain in the

previous 24-hours) samples were successfully collected within any rolling

30-day period 7 times in 2014, 4 times in 2015, 0 times in 2016, and 1

time in 2017. These periods were not well distributed throughout the

swimming season, which emphasizes the need to specifically target 5

base flow events for each 30-day period. An excursion of the 5-day

geometric mean criterion for fecal coliform of 200/100 ml occurred for the

periods of 8/19/2014 – 9/16/2014 and 8/26/2014 – 9/23/2014 at Site 6;

8/19/2014 – 9/16/2014, 8/26/2014 – 9/23/2014, and 5/26/2015 –

2-23

6/23/2015 at Site 7; and 8/19/2014 – 9/16/2014 at Site 8. A total of 6 30-

day geometric mean excursions were documented at three sites.

Exceedances of the fecal coliform criterion, “no more than 10% of the total

samples taken during a 30-day period may exceed 400 per 100 ml,”

occurred at most sites throughout the four years of data. No exceedances

occurred at Site 1. Site 2 and Site 7 had the most exceedances and had

exceedances for each year (Table 1). In addition to base flow events,

multiple wet weather events (> 0.25” within 24-hours) were sampled 7 – 9

times through the swimming season each year. The highest maximum and

mean values were documented at Site 6 and Site 7 (Table 2, Figure 1).

Table 1. Number of exceedances for fecal coliform of no more than 10% >

400/100ml for rolling 30-day periods at sites for specific years.

Year Site 1 Site 2 Site 3 Site 4 Site 6 Site 7 Site 8

2014 0 5 0 2 4 5 2

2015 0 3 0 2 3 5 3

2016 0 1 1 0 0 5 0

2017 0 1 1 1 1 1 1

Table 2. Wet weather event yearly summaries (fecal coliforms/100 ml)

Year Statistic Site 1 Site 2 Site 3 Site 4 Site 6 Site 7 Site 8

2014

Min 18 18 45 68 18 170 78

Max 220 1100 330 1300 2400 7900 3500

Mean 88.5 336.8 154.2 595.4 605.1 1812.8 894

2015

Min 55.6 42.5 22.1 410.6 290.9 613.1 365.4

Max 782.5 776.5 2053 2419.6 3065.5 7068 3635

Mean 372.5 351.2 772.4 1214.0 1037.6 2793.1 1324.0

2016

Min 7.5 54 21.3 19.5 36.8 1301.5 91

Max 365.4 461.1 1046.2 1046.2 2419.6 12098 2442

Mean 155.4 224.9 254.0 407.2 870.9 3083.9 1017.3

2017

Min 55.6 15 40.4 111.9 88 111.2 85.7

Max 1827 3448 2735.5 2735.5 3065.5 2897 2586

Mean 414.6 713.5 594.7 603.4 987.2 1264.4 678.2 * Bold values indicate the greatest maximum and mean values for a given year

2-25

USEPA. 2011. CSO post construction compliance monitoring guidance. EPA-833-K-11-

001. U.S. Environmental Protection Agency, Office of Wastewater Management,

Washington, DC.

3-2

3.1 WADEABLE RIFFLE-RUN STREAM MACROINVERTEBRATE DATA

COLLECTION PROTOCOL

3-3

Prepared by:

Brian Chalfant






2013

Edited by:

Dustin Shull






2017

3-4

INTRODUCTION

USEPA’s Rapid Bioassessment Protocols for Use in Streams and Wadeable Rivers

(Barbour et al.1999) describes two general approaches to collecting stream

macroinvertebrate communities for water quality assessment purposes. These

approaches are the “single, most productive habitat” approach, described here, and

the “multihabitat” approach, later described in Chapter 3.3, Wadeable Multihabitat

Stream Macroinvertebrate Collection Protocol (Pulket 2017). The single, most

productive habitat approach is typically used to assess streams where cobble

substrate and riffle/run habitat is predominant (generally referred to as high gradient

freestone streams). This protocol identifies appropriate field collection procedures

needed to evaluate macroinvertebrate communities in wadeable freestone streams

where riffle-run habitat is sufficiently abundant and is a modification of the USEPA

Rapid Bioassessment Protocols (Barbour et al. 1999). Samples can be collected year-

round; however, certain periods of the year are required depending on the purpose of

the study. Refer to Chapter 2.1, Wadeable Freestone Riffle-Run Stream

Macroinvertebrate Assessment Method (Shull 2017b) of the Assessment Book for

more information on collection results and season.

MACROINVERTEBRATE FIELD COLLECTION

A benthic macroinvertebrate sample collection begins with delineating a 100-meter

reach along the stream where the best available representation of riffle-run habitat

exists for the stream segment of interest. Within the reach, six one-minute (each kick

must be at least 45 to 60 seconds in duration) kicks are conducted immediately

upstream of a non-truncated D-framed net with 500 µm mesh (Figure 1). Truncated nets

are shorter with less meshed surface area and are not recommended for this collection

protocol.

Each kick should disturb approximately 1 m2 immediately upstream of the net to an

approximate depth of 10 cm, as substrate allows. During each kick, the net should be

held stationary. Kicks are completed in a downstream-to-upstream direction to avoid

disturbing the upstream portions of the targeted reach. Collectors must ensure kicks are

distributed throughout the 100-meter reach and are representative of the variety of riffle-

run habitats present (e.g., slow-flowing, shallow riffles and fast-flowing, deeper riffles). It

is important to note that riffle-run habitat differences will be respective of the stream

segment being sampled. For example, a slow-flowing deep riffle will be different in a

small stream than in a larger stream. Kicks must also be conducted throughout the

width of the stream to include the left-descending, middle, and right-descending areas.

The only exception to this is if a specific area of the stream is being targeted for a

3-5

reason. If that is the case, then the collector must clearly document that the sample is

not a complete composite and state the purpose for collection on the sample form.

Figure 1. A non-truncated D-framed net with 500 µm mesh.

When collecting same day replicate samples within the same reach, it is important not

to disturb the same areas of the stream with both sets of kicks. It is also important to be

cognizant of macroinvertebrate drift after kicking the first sample. For example, a kick

during the replicate collection may be very close and immediately downstream of an

original sample kick without being in the same location. Disturbance from the original

kick just upstream of a replicate kick may have disturbed macroinvertebrates and skew

the replicate sample. An example of good kick selection and incorrect kick selection is

illustrated in Figure 2. It is recommended that a replicate sample be collected after

every 10 regular samples for QA/QC purposes.

3-6

Figure 2. Blue rectangles illustrate a 100-meter reach within a stream, dark blue

ellipses show the initial set of kicks during sample collection, and yellow ellipses show

the replicate set of kicks during sample collection. (A) Correct sample collection with

replicate collection; all kicks are collected within 100 meters and across the entire width

of the stream, replicate kicks are not collected in the same location, and replicate kicks

are not placed in close proximity, directly downstream of any other kicks. (B) Incorrect

sample collection (with replicate); not all kicks are collected within 100 meters and

across the width of the stream, replicate kicks are collected in the same location, and in

close proximity, directly downstream of other kicks.

Once collected from the stream, each of the six kicks are composited into one sample

jar (or as few samples jars as possible). Each jar is labeled with the date, time, collector,

collectively referred to as a station ID (yyyymmdd-HHMM-Collector) and location. If

more than one jar is needed then each jar will also be labeled with a number (e.g., 1 of

2, 2 of 2). It is recommended that paper tags labeled using indelible ink or pencil be

placed inside the sample jar(s) as well. Care must be taken to minimize damage to the

collected organisms when compositing the materials. For this reason, only larger rocks,

3-7

detritus, and other debris are rinsed and removed before the sample is preserved.

Buckets and 500 µm mesh sieves are permitted when compositing kicks; however,

swirling the composited material in a bucket filled with water to reduce the amount of

debris (elutriation) is not recommended as this action can unnecessarily damage fragile

organisms, or allow for heavier organisms to be discarded unintentionally. Composited

samples are preserved with at least 70% ethanol (95% ethanol is preferred) in the field.

Collectors must ensure that the percentage of ethanol is high enough to properly

preserve the sample and that there is ample space left in the sample jar to properly

preserve organisms. Composited samples are then transported back to the laboratory

for processing and all field information is recorded on a macroinvertebrate field data

sheet (Appendix A.1) or mobile data collection application.

ADDITIONAL DATA

During a DEP riffle-run macroinvertebrate survey, field meter (temperature, specific

conductance, pH, dissolved oxygen, and turbidity if available), water chemistry, and

habitat assessment data should also be collected. Recommended Standard Analysis

Codes (SAC) for riffle-run macroinvertebrate collections are SAC 612 or SAC 18 (for

special protection). For more information on water chemistry collection and SACs, refer

to Chapter 4.2, Discrete Water Chemistry Data Collection Protocol (Shull 2017a).

Because riffle-run surveys are designed for high gradient streams, the high gradient

habitat assessment form should be used. The DEP riffle/run prevalent habitat form can

be found in Appendices C.1 and C.2.

LITERATURE CITED

Barbour, M.T., J. Gerritsen, B. D. Snyder, and J. B. Stribling. 1999. Rapid

bioassessment protocols for use in streams and wadeable rivers: Periphyton,

benthic macroinvertebrates and fish. 2nd edition. EPA 841-B-99-002. U.S.

Environmental Protection Agency, Office of Water, Washington, D.C.

Pulket, M. (editor). 2017. Wadeable multihabitat stream macroinvertebrate data

collection protocol. Chapter 3.3, pages 14–19 in M. J. Lookenbill, and R.



Pennsylvania.

Shull, D. R. 2017a. Discrete water chemistry data collection protocol. Chapter 4.2,

pages 9–24 in M. J. Lookenbill, and R. Whiteash (editors). Water quality



3-8

Shull, D. R. (editor). 2017b. Wadeable freestone riffle-run stream macroinvertebrate

assessment method. Chapter 2.1, pages 2–24 in D. R. Shull, and R. Whiteash



3-9

3.2 WADEABLE LIMESTONE STREAM MACROINVERTEBRATE DATA

COLLECTION PROTOCOL

3-10

Prepared by:

William Botts






2009

Edited by:

Amy Williams






2017

3-11

INTRODUCTION

Limestone streams are unique aquatic systems that are typically very popular for

recreational fishing. To properly protect the ecological integrity of limestone streams,

they must be assessed correctly. This data collection protocol is intended to aid in

assessing the aquatic life uses of Pennsylvania’s wadable limestone streams. In the

case of these limestone stream assessments, the following field procedures will apply.

A stream must meet certain criteria for this protocol to be applicable for assessments

(Table 1). Limestone streams will often display low-gradient flow conditions and are

almost completely void of clearly defined freestone habitat beds.

Table 1. Limestone streams definition criteria

Parameter Criterion Explanation

Alkalinity Minimum 140 mg/l Stream must maintain high

alkalinity throughout the year

Temperature 40 to 65°F 4 to 18°C

Constant temperatures are very

important. Check to see if stream

is ice-free in the winter.

Known Influences

Stream originates from

limestone springs or very

strongly influenced by

limestone springs.

N/A

Drainage Area Maximum 20 sq. miles

There may be exception to this

parameter if all other criteria are

met.

Water Use Definition of Cold Water

Fishes (CWF)

Meets the definition of a CWF

stream in Chapter 93.

FIELD SAMPLING CONSIDERATIONS

Net Mesh & Sieve

Many state water quality programs, federal agencies (e.g., USEPA, USGS), and other

water quality monitoring organizations use net sampling devices with 500μm mesh nets.

Due to the natural conditions of limestone streams, 500μm mesh size quickly clogs,

preventing macroinvertebrates and vegetation from entering the net resulting in a poor

sample. To ensure an accurate assessment, 800 - 900μm net mesh must be used to

collect samples. A standard 500μm mesh sieve should be used to process samples.

3-12

Semi-Quantitative Procedure

The DEP Rapid Bioassessment Protocol (RBP) is a modification of the USEPA RBP III

(Plafkin et al. 1989). Modifications include: 1) the use of a D-frame net for the collection

of the riffle-run samples, 2) different laboratory sorting procedures, 3) elimination of the

CPOM (coarse particulate organic matter) sampling, and 4) metrics substitutions. Unlike

USEPA’s RBP III methodology, no field sorting is done. Only larger rocks, detritus, and

other debris are rinsed and removed while in the field before the sample is preserved.

While USEPA’s RBP III method was designed to compare impacted waters to reference

conditions (cause and effect approach), the DEP-RBP modifications were designed for

unimpacted waters, as well as impacted waters.

FIELD PROCEDURES

Benthic Macroinvertebrate Sampling

The handheld D-frame sampler consists of a bag net attached to a half-circle (“D”

shaped) frame that is 1 ft. wide. The net is employed by one person facing downstream

and holding the net firmly on the stream bottom. One “D-frame effort” is defined as:

vigorous kicks in an approximate area of 1m2 (1 x 1 m) immediately upstream of the net

to a depth of 10cm (approximately 4 inches, as the substrate allows) for approximately

one minute. All benthic dislodgement and substrate scrubbing should be done by kicks

only. Substrate handling should be limited to the removal of large rocks or debris (as

needed) with no hand washing. Since the width of the kick area is wider than the net

opening, net placement is critical to assure all kicked material flows toward the net.

Avoiding areas with crosscurrents, the substrate material from within the 1 m2 area

should be kicked toward the center of the square meter area.

The purpose of the standardized DEP-RBP sampling procedure is to obtain

representative macroinvertebrate fauna from comparable stations. The DEP-RBP

assumes the riffle/run to be the most productive habitat. Riffle/run habitats are sampled

using the D-frame net procedure described above. For limestone stream surveys, two

D-frame efforts (kicks) are collected - one from an area of fast velocity and one from an

area of slower velocity within the same riffle. Limestone streams have low gradient often

making it difficult to locate well developed riffles. If there are no riffles in the sample

area, a run, or the best rock substrate available is sampled. The two kicks are then

composited into one sample jar (or more as necessary). Each jar is labeled with the

date, time, collector, collectively referred to as a station ID (yyyymmdd-HHMM-

Collector) and location. If more than one jar is needed then each jar will also be labeled

with a number (e.g., 1 of 2, 2 of 2). It is recommended that paper tags labeled using

indelible ink or pencil be placed inside the sample jar(s) as well. Care must be taken to

minimize damage to the collected organisms when compositing the materials. It is

3-13

recommended that the benthic material be placed in a bucket filled with water to

facilitate gentle stirring and mixing as not to damage fragile organisms. Because

limestone samples are very abundant in organic material, the sample is preserved with

at least 70% ethanol (95% ethanol is preferred). Composited samples are then

transported back to the laboratory for processing and all field information is recorded on

a macroinvertebrate field data sheet (Appendix A.1) or mobile data collection

application.

Sample Collection Period

Samples must be collected from January through May. Limestone streams have a low

number of sensitive taxa and only a few of these taxa are generally found in large

numbers. One very important sensitive taxon is Ephemerella. A good population of

Ephemerella generally indicates better water quality. The three species of Ephemerella:

invaria (rotunda) and dorothea found in Pennsylvania limestone streams emerge in May

and June and are normally difficult or impossible to collect after emergence and until

nymphs mature. Collecting samples from January through May ensures these very

important ecological indicator taxa will not be missed.

ADDITIONAL DATA

During a DEP limestone macroinvertebrate survey, field meter (temperature, specific


habitat assessment data should also be collected. Recommended Standard Analysis

Codes (SAC) for limestone macroinvertebrate collections are SAC 612 or SAC 18 (for

special protection). For more information on water chemistry collection and SACs, refer

to Chapter 4.2, Discrete Water Chemistry Data Collection Protocol (Shull 2017). The

Riffle/Run Prevalence Habitat assessment form should be used for limestone

macroinvertebrate collections. This form can be found in Appendices C.1 and C.2.

LITERATURE CITED

Plafkin, J. L, M. T. Barbour, K. D. Porter, S. K. Gross, and R. M. Hughes. 1989. Rapid

bioassessment protocols for use in rivers and streams: Benthic

macroinvertebrates and fish. EPA/440/4-89-001. U.S. Environmental Protection

Agency, Office of Water, Washington, D.C.

Shull, D. R. 2017. Discrete water chemistry data collection protocol. Chapter 4.2, pages




3-14

3.3 WADEABLE MULTIHABITAT STREAM MACROINVERTEBRATE DATA

COLLECTION PROTOCOL

3-15

Prepared by:

Charles McGarrell and Molly Pulket






2007

Edited by:

Molly Pulket






2017

3-16

INTRODUCTION

USEPA’s Rapid Bioassessment Protocols for Use in Streams and Wadeable Rivers

(Barbour et al.1999) describes two general approaches to assessing stream

macroinvertebrate communities. These approaches are the “single, most productive

habitat” approach, previously described in Chapter 3.1, Wadeable Riffle-Run Stream

Macroinvertebrate Data Collection Protocol (Shull 2017b), and the “multihabitat”

approach, described here. The multihabitat approach involves sampling a variety of

habitat types instead of sampling a single habitat, such as cobble substrate in riffles

and/or runs. This data collection protocol identifies appropriate field collection

procedures needed to evaluate macroinvertebrate communities in wadeable low-

gradient Pennsylvania streams.

A low-gradient waterway in Pennsylvania is defined as having pool/glide channel

morphology and naturally lacking riffles. There should be no riffles within 300

consecutive meters of the sampling reach. Cobble outcrops extending less than 1/8 of

the stream width are not considered riffles. In addition, small areas with slightly

increased velocities and gravel substrates are not considered riffles. Besides lack of

riffle/run sequences, low-gradient streams are characterized by a gradient less than

0.5%, substrates of fine sediment or infrequent aggregations of coarse sediments, and

an aggregate of the five habitat types as described in Table 1. Moderate or high

gradient stream segments that have been influenced by impoundments or beaver

dams are not appropriate sites for the multihabitat field sampling protocol.

MACROINVERTEBRATE FIELD COLLECTION

Aquatic macroinvertebrate samples are collected using a multihabitat sample collection

procedure modified from that described in Barbour et al. (1999). Sampling should occur

during the months of November through May. A 100-meter length of stream is

determined and set as the sample reach. The investigator then identifies which low-

gradient habitat types are present within the sample reach. Table 1 describes the five

habitat types and explains the different sampling techniques for each habitat. Sampling

consists of 10 D-frame net jabs, 2 in each of the five habitat types or distributed

proportionally in the available habitats. A minimum surface area of approximately 0.46

m2 is required for a given habitat type to be sampled. If the total number of jabs (10) is

not evenly divisible by the number of habitat types present, the remaining jab(s) are

distributed among the most abundant habitat type(s) in the reach. Each jab consists of a

30-inch-long sweep of a 0.3-meter wide area, using a 500-micron mesh D-frame dip

net. The jabs are composited while the investigator works progressively upstream from

the first collection site. After the 10 jabs are completed, all jabs are combined into

3-17

sampling jars and preserved with at least 70% ethanol (95% ethanol is preferred) in the

field. Each jar is labeled with the date, time, collector (collectively referred to as a station

ID (yyyymmdd-HHMM-Collector)), and location. If more than one jar is needed, then

each jar will also be labeled with a number (e.g., 1 of 2, 2 of 2). Jars should only be

filled 2/3 full of debris to allow for adequate sample preservation (ethanol should fill jar

completely). If a lot of debris or large pieces were collected, the debris can be carefully

rinsed in buckets, inspected for organisms, and then discarded. The rinse water is

poured through a 500 µm mesh sieve to retain the macroinvertebrates before being

discarded. The net, sieve, and bucket should be thoroughly inspected for

macroinvertebrates. Composited samples are then transported back to the laboratory

for processing and all field information is recorded on the macroinvertebrate field data

sheet (Appendix A.1) or mobile data collection application.

3-18

Table 1. Stream Habitat Types and Field Sampling Techniques

Habitat Type Description Sample Technique

Cobble/ Gravel Substrate

Stream bottom areas consisting of mixed gravel and larger substrate particles; Cobble/gravel substrates are typically

located in relatively fast-flowing, “erosional” areas of the stream channel

Macroinvertebrates are collected by placing the net on the substrate near the downstream end of an area of gravel or larger substrate particles and simultaneously pushing down on the net while pulling it in an upstream direction with adequate force to dislodge substrate materials and

the aquatic macroinvertebrate fauna associated with these materials; Large stones and organic matter

contained in the net are discarded after they are carefully inspected for the presence of attached organisms which

are removed and retained with the remainder of the sample; One jab consists of passing the net over

approximately 30 inches of substrate.

Snag

Snag habitat consists of submerged sticks, branches, and other woody debris that appears to have been submerged long enough to be adequately colonized by

aquatic macroinvertebrates; Preferred snags for sampling include small to medium-sized sticks and branches (preferably < ~4 inches

in diameter) that have accumulated a substantial amount of organic matter (twigs, leaves, uprooted aquatic macrophytes, etc.)

that is colonized by aquatic macroinvertebrates.

When possible, the net is to be placed immediately downstream of the snag, in either the water column or on

the stream bottom, in an area where water is flowing through the snag at a moderate velocity; The snag is then kicked in a manner such that aquatic macroinvertebrates

and organic matter are dislodged from the snag and carried by the current into the net; If the snag cannot be

kicked, than it is sampled by jabbing the net into a downstream area of the snag and moving it in an

upstream direction with enough force to dislodge and capture aquatic macroinvertebrates that have colonized

the snag; One jab equals disturbing and capturing organisms from an area of ~0.23 m2 (12” x 30”)

Coarse Particulate

Organic Matter (CPOM)

Coarse particulate organic matter (CPOM) consists of a mix of plant parts (leaves, bark, twigs, seeds, etc.) that have accumulated on the stream bottom in “depositional” areas of

the stream channel; In situations where there is substantial variability in the

composition of CPOM deposits within a given sample reach (e.g., deposits

consisting primarily of white pine needles and other deposits consisting primarily of hardwood tree leaves), a variety of CPOM deposits are sampled; However, leaf packs in higher-velocity (“erosional”) areas of the channel are not included in CPOM samples

CPOM deposits are sampled by lightly passing the net along a 30-inch long path through the accumulated

organic material to collect the material and its associated aquatic macroinvertebrate fauna; When CPOM deposits are extensive, only the upper portion of the accumulated organic matter is collected to ensure that the collected

material is from the aerobic zone

Submerged Aquatic

Vegetation (SAV)

Submerged aquatic vegetation (SAV) habitat consists of rooted aquatic macrophytes

SAV is sampled by drawing the net in an upstream direction along a 30-inch long path through the

vegetation; Efforts should be made to avoid collecting stream bottom sediments and organisms when sampling

SAV areas.

Sand/Fine Sediment

Sand/fine sediment habitat includes stream bottom areas that are composed primarily of

sand, silt, and/or clay.

Sand/fine sediment areas are sampled by bumping or tapping the net along the surface of the substrate while slowly drawing the net in an upstream direction along a 30-inch long path of stream bottom; Efforts should be

made to minimize the amount of debris collected in the net by penetrating only the upper-most layer of sand/silt

deposits; Excess sand and silt are removed from the sample by repeatedly dipping the net into the water column and lifting it out of the stream to remove fine

sediment from the sample

3-19

ADDITIONAL DATA

During a DEP multihabitat macroinvertebrate survey, field meter (temperature, specific


habitat assessment data should also be collected. Recommended SAC for multihabitat

collections are SAC 612, SAC 87, and SAC 18 (for special protection). For more

information on water chemistry collection and SACs, refer to Chapter 4.2, Discrete

Water Chemistry Data Collection Protocol (Shull 2017a). Because multihabitat surveys

are designed for low gradient streams, the low gradient habitat assessment form should

be used. The DEP low gradient habitat form can be found in Appendix C.3.

LITERATURE CITED

Barbour, M. T., J. Gerritsen, B. D. Snyder, and J. B. Stribling. 1999. Rapid








Shull, D. R. (editor). 2017. Wadeable riffle-run stream macroinvertebrate data collection

protocol. Chapter 3.1, pages 2–8 in M. J. Lookenbill, and R. Whiteash (editors).

Water quality monitoring protocols for streams and rivers. Pennsylvania


3-20

3.4 SEMI-WADEABLE LARGE RIVER MACROINVERTEBRATE DATA

COLLECTION PROTOCOL

3-21

Prepared by:

Dustin Shull






2017

3-22

INTRODUCTION

Collection of macroinvertebrate data in large semi-wadeable rivers can be a daunting

and complex process. To appropriately collect biological community data in large rivers

and to increase efficiency, DEP separates large rivers into two categories, semi-

wadeable and non-wadeable. Semi-wadeable rivers are defined as predominantly free-

flowing systems with drainage areas > 1,000 mi2 and have physical characteristics that

allow for riffle and run sections to occur with relative frequency. These river systems

tend to lack a well-defined and navigable U-shaped channel for any significant distance

and frequently present difficulties for both wadeable and non-wadeable

macroinvertebrate data collection protocols. In addition, large semi-wadeable rivers

potentially have differences in water quality across their width. Well over half of the

large rivers within Pennsylvania are considered semi-wadeable (Figure 1). Therefore, it

is important that a standardized protocol be established for the collection of physical,

chemical, biological data in these river systems.

Figure 1. Semi-wadeable and non-wadeable rivers throughout Pennsylvania.

The goal of this document is to lay the framework for how DEP intends on collecting

macroinvertebrate and supplementary data for making Aquatic Life Use (ALU)

determinations in large semi-wadeable rivers within the boundaries of Pennsylvania.

3-23

Working on, in, and around large semi-wadeable rivers can be challenging and

dangerous. This discussion seeks to provide guidance on when and how to collect

these data, as well as important safety considerations that must be considered. Making

accurate and defensible assessment decisions requires both enough data types (e.g.,

physical, chemical, and biological) and a specificity of those data within a water

influence (zone) and season.

SAFETY PROCEDURES

Appropriate ALU assessment begins with data collection that follows proper guidelines

and safety procedures. Safety is of utmost concern when working in and on large semi-

wadeable rivers. Individuals that lack experience and education related to navigating

these waterbodies put themselves and others at risk of injury or loss of life. In most, if

not all cases, some form of watercraft is required for this sampling protocol. Whether a

kayak, canoe, or powered boat is used, DEP recommends – and in most cases PFBC

requires by law – that individuals using watercrafts take boating courses and obtain a

boating safety certificate. However, whether laws apply to certain individuals or

watercraft or not, DEP strongly recommends that all individuals regardless of age,

experience, or navigation procedure obtain and keep a boating safety certification on

them during semi-wadeable surveys. More information on boating courses and safety

certifications are available here:

http://www.fishandboat.com/Boat/BoatingCourses/Pages/default.aspx

Large semi-wadeable rivers present difficulties for both wadeable and non-wadeable

water navigation. Certain locations on rivers such as the Susquehanna River around

Harrisburg, PA may be fully or partially wadeable during certain times of year, but there

are also highly variable currents and drop-offs that are dangerous to wading collectors.

Therefore, personal floatation devices (PFD) should always be worn. However, PFDs

should not be used to facilitate wading or navigating the waterbody. Navigation using a

watercraft can be equally if not more challenging than wading. Many boats that frequent

the large semi-wadeable rivers such as the Susquehanna River are outfitted with

modified jet propulsion to reduce the risk of serious damage associated with impacting

the streambed with the lower unit. Therefore, consideration for modified equipment may

be needed to conduct this protocol. In addition, riffles and bedrock substrate often

dominate these rivers regardless of size. Consequently, DEP recommends that at least

two individuals work together during these surveys. This is critical for many safety

reasons, particularly important when a powered watercraft is underway. While in motion,

DEP recommends that the individual not operating the powered watercraft is actively

looking for obstacles on and just under the surface. This individual should be actively

and clearly communicating obstacles to the operator. This is best done with the use of

3-24

hand signals that are agreed upon between collectors prior to navigating the water. All

of this is not to say that macroinvertebrate sampling in large semi-wadeable rivers is

overly risky. In fact, with the proper training and experience this sampling protocol can

be conducted efficiently and safely using limited resources, and without restrictions

based on the collector’s physical stature.

Inherent in these recommendations is the understanding that collectors must be well

informed of the unique characteristics of each river they intend to sample and be aware

of current flow and weather conditions. Equipment considerations may be different

between similar sized large semi-wadeable rivers, and certainly different when visiting

multiple semi-wadeable rivers of different sizes. For instance, boat selection and safety

equipment lists would be different depending on whether the collector is visiting a 1,000

mi2 river or 10,000 mi2 river. These differing characteristics require thorough planning

before going into the field, so resources are not wasted due to a lack of preparation. It is

also imperative that collectors have access to and continually check river flow

conditions. There will be situations when weather in one part of Pennsylvania may be

optimal for sampling a site, but weather in another part of Pennsylvania – two or three

days before the sampling period – created conditions that reduces the ability to collect.

Analysis of flow conditions at all available points upstream of the intended sampling

locations are highly recommended. Commonly used discharge data are available here:

https://waterdata.usgs.gov/pa/nwis/current.

WATER QUALITY DATA COLLECTION

Cross-section surveys and water chemistry samples must be collected before

macroinvertebrate sampling. This is needed to determine where the macroinvertebrate

sample(s) are collected. All cross-section surveys are conducted using a clean and

calibrated field meter that collects water temperature, specific conductance, dissolved

oxygen, pH, and – preferably – turbidity (see Chapter 4). Transects (discrete points

within the cross-section survey) are taken at regular intervals (each point representing

approximately 5-10% of the total wetted width) across the width of the river. This is the

least amount of information required to determine if major water quality influences exist

at the sampling location. A major water quality influence that shows a 10% difference or

greater in any field meter parameter necessitates a unique water chemistry and

macroinvertebrate sample be collected. It is often helpful to create a drawing or map of

the site with transect points depicted to help delimit the locations of water influence

transitions across the width of the river (Figure 2). When new major water influences are

identified, additional transects and water chemistry samples will be collected on

upstream reaches to identify and characterize the source of variable water quality

across the width of the targeted reach of river. Cross-section surveys and water

3-25

chemistries will also be collected downstream, if possible, to determine the extent that

the water influences do not mix. Multiple water chemistry samples over a period are

recommended to provide additional information for assessments. Cross-section surveys

and water chemistry data collected through the season prior to macroinvertebrate

collection provides the most utility for complete and accurate assessments, but routine

year-round data collection has also proven useful for ALU assessments and method

development efforts. Cross-section surveys and water chemistry data collection prior to

macroinvertebrate collections also provides opportunities to complete a thorough

reconnaissance of the targeted semi-wadable river.

Figure 2. Example of cross-section data collection where data are collected at

established transect points over time. In this example, the cross-section survey data

identify three unique major water influences that create the three zones where

macroinvertebrate, chemical, and physical data should be collected.

There are three water chemistry collection protocols that are acceptable when collecting

chemical data on semi-wadeable rivers: fixed-width sampling, isokinetic sampling

(USGS 2006), and discrete mid-channel/mid-depth sampling (see Chapter 4.2, Discrete

Water Chemistry Data Collection Protocol (Shull 2017a)). Fixed width and isokinetic

3-26

sampling is useful for characterizing water chemistry for the entire width of the surface

water, while discrete samples are useful for characterizing unique water quality

influences. Water chemistry characterization is also driven by the need to identify

sources and causes of impairment. Water chemistries should include a comprehensive

list of constituents including total and dissolved nitrogen and phosphorus species, total

metals, and ions. For water chemistry collection, DEP routinely uses SAC 612 for semi-

wadeable rivers. In addition, constituents such as dissolved metals, pesticides,

hormones, wastewater compounds, and pharmaceuticals are added if necessary. For

more information on water chemistry collection and SACs, refer to Chapter 4.2, Discrete

Water Chemistry Data Collection Protocol (Shull 2017a).

MACROINVERTEBRATE COLLECTION

Semi-wadeable large river macroinvertebrate samples are collected using 6D-200 field

collection described in the Wadeable Riffle-Run Stream Macroinvertebrate Data

Collection Protocol (Shull 2017b). Briefly, this consists of 6 one-minute kicks evenly

distributed across the entire width of the waterbody using a 500 µm mesh D-frame net.

With the transect procedure modification, additional 6D-200 samples are collected in

accordance with the number of major water quality influences discovered during water

quality transect data collection. For instance, if three distinctly different water influences

were discovered, then three 6D-200 samples are collected, with each sample

composited across the entire width of the respective water influence. Jars are labeled

with the date, time, collector, collectively referred to as a station ID (yyyymmdd-HHMM-

Collector), and location description. If more than one jar is needed then each jar will

also be labeled with a number (e.g., 1 of 2, 2 of 2). Samples are preserved with at least

70% ethanol (95% ethanol is preferred). Once field collections are complete, samples

are taken to the laboratory for analysis. Macroinvertebrate processing and identification

is conducted using standard DEP protocols, which are also described in detail in

Chapter 3.6, Macroinvertebrate Laboratory Subsampling and Identification Protocol

(Brickner 2020).

Macroinvertebrate samples collected at or above median flow conditions can cause a

lower confidence in the results. Therefore, if collectors are not confident that the

sampling was representative of the zone (e.g., collector could not collect the 6 kicks

relatively evenly across the entire width of the influence) or they have collected in an

appropriate habitat type, then samples should not be used for assessments. The

responsibility for this is decision is left to the most experienced individual in the field

during the sampling event. Individuals that are experienced with working in large rivers

quickly realize when an inappropriate sample has been collected and will either discard

3-27

the sample or annotate their concerns in the comments section of the macroinvertebrate

data collection field form (Appendix A.1) or mobile data entry application.

ADDITIONAL DATA

Additional field meter (temperature, specific conductance, pH, dissolved oxygen, and

turbidity if available) and water chemistry data does not need to be collected if the

cross-sectional surveys and chemistry collection were completed as described above. A

habitat assessment should also be collected for every macroinvertebrate collection.

Because semi-wadeable large river surveys target riffle-run habitat, the DEP Riffle-Run

Prevalence Habitat Form should be used. This form is found in Appendix C.1 and C.2.

LITERATURE CITED

Brickner, M. (editor). 2020. Macroinvertebrate laboratory subsampling and identification

protocol. Chapter 3.6, pages 32–42 in M. J. Lookenbill and R. Whiteash (editors).

Water quality monitoring protocols for streams and rivers. Pennsylvania






Shull, D. R. (editor). 2017b. Wadeable riffle-run stream macroinvertebrate data

collection protocol. Chapter 3.1, pages 2–8 in M. J. Lookenbill, and R. Whiteash

(editors). Water quality monitoring protocols for streams and rivers. Pennsylvania


USGS. 2006. National field manual for the collection of water quality data. Chapter 4.1

in Surface water sampling: Collection methods at flowing-water and still-water

sites. U.S. Geological Survey, Water Resources Office of Water Quality, Reston,

VA. (Available online at:

https://water.usgs.gov/owq/FieldManual/Chapter4/pdf/Chap4 v2.pdf.)

3-28

3.5 QUALITATIVE BENTHIC MACROINVERTEBRATE DATA COLLECTION

PROTOCOL

3-29

Prepared by:

Tony Shaw






2006

Edited by:

Josh Lookenbill






2017

3-30

INTRODUCTION

While DEP primarily relies on semi-quantitative benthic macroinvertebrate data

collection protocols described previously in this chapter, qualitative data collection

protocols could also be considered. Qualitative macroinvertebrate data collection

protocols may be appropriate for cause and effect surveys and point of first use

surveys; however, this collection protocol is no longer used for ALU determinations.

The type of sampling gear used is dependent on waterbody type, available habitat, and

other site-specific conditions. The recommended gear for sampling wadeable streams is

a 1m2, flexible kick-screen, or a non-truncated D-framed net. The type of gear,

dimensions, and mesh size must be reported for all data collections. When more than

one gear type is used, the results must be recorded separately. Physical stream

variables should be matched as closely as possible when comparing reference stations

to candidate/impacted stations during location placement. Gear-type should also match

between stations within the same survey. Matching these variables helps minimize or

eliminate the effects of compounding variability.

KICK-SCREEN

A common qualitative sampling protocol uses a simple hand-held kick-screen. This

device is designed to be used by two persons. However, with experience, it may be

used by one person. The kick-screen is constructed with a 1x1meter piece of net

material fastened to two dowel handles.

Facing up stream, one person places the net in the stream with the bottom edge of the

net held firmly against the streambed. An assistant then vigorously kicks the substrate

within a 1x1m area immediately upstream of the net to a depth of 7-10cm, as substrate

allows. The functional depth sampled may vary due to ease of disturbance as

influenced by substrate embeddedness. The amount of effort expended in collecting

each sample should be approximately equivalent to make valid comparisons.

As many as four screens are collected at each site or until no new taxa are collected.

Initial sampling should target riffle/run habitat. Collection in additional habitats to

generate a more complete taxa list can be conducted at the discretion of the

investigator.

Station data will be recorded on the macroinvertebrate data collection forms

(Appendices A.1 and A.2) and habitat data will be recorded on the appropriate habitat

data collection forms (Appendices C.1, C.2, or C.3) in this book.

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D-FRAMED NET

DEP has standardized a non-truncated D-framed net with 500µm mesh for all semi-

quantitative benthic macroinvertebrate data collections in freestone streams and rivers,

however it may also be appropriate to use this gear for qualitative collections.

Qualitative D-framed data collection could be employed where the width of the targeted

waterbody is less than 1 meter, or the targeted reach is small with very little available

habitat.

In wadable, flowing waters with available riffle/run habitat, a D-framed net can be

employed using a “single, most productive habitat” approach, previously described in

this chapter. Facing upstream, each one-minute (each kick must be at least 45 to 60

seconds in duration) kick is conducted immediately upstream of a non-truncated D-

framed net. Each kick should disturb approximately 1m2 immediately upstream of the

net to an approximate depth of 10 cm, as substrate allows. During each kick, the net

should be held stationary. With semi-quantitative protocols a prescribed, a standardized

effort (number of kicks) is required. Qualitatively this gear should be used until no new

taxa are collected. Kicks are completed in a downstream-to-upstream direction to avoid

disturbing the other targeted portions of reach before kicks are conducted in that area.

Collectors must ensure kicks are representative of the variety of riffle-run habitats

present (e.g., slow-flowing, shallow riffles and fast-flowing, deeper riffles). It is important

to note that riffle-run habitat differences will be respective of the stream segment being

sampled. For example, a slow-flowing deep riffle will be different in a small stream than

in a larger stream. Kicks must also be conducted throughout the width of the stream to

include the left-descending, middle, and right-descending areas. The only exception to

this is if a specific area of the stream is being targeted for a reason. If that is the case,

then the collector must clearly document that the sample is not a complete composite

and state the purpose for collection on the sample form.

In wadable, low-gradient waters without riffle/run habitat the “multihabitat” approach

may be appropriate. The multihabitat approach involves sampling a variety of habitat

types instead of sampling a single habitat, such as cobble substrate in riffles and/or

runs. The investigator first identifies which low-gradient habitat types are present within

the sample reach. The five habitat types and the different sampling techniques are

described in Chapter 3.3, Wadeable Multihabitat Stream Macroinvertebrate Data

Collection Protocol (Pulket 2017). Qualitatively this gear should be used until no new

taxa are collected. Sampling consists of D-frame net jabs, in each of the available

habitat types. A minimum surface area of approximately 0.46 m2 is required for a given

habitat type to be sampled. Each jab consists of a 30-inch-long sweep of a 0.3-meter

3-32

wide area, using a 500-micron mesh D-frame dip net. Jabs are completed in a

downstream-to-upstream direction to avoid disturbing the other targeted portions of

reach before jabs are conducted in that area.

ADDITIONAL DATA

Field meter (temperature, specific conductance, pH, dissolved oxygen, and turbidity if

available), water chemistry, and habitat assessment data should also be collected.

Station data will be recorded on the macroinvertebrate data collection forms

(Appendices A.1 and A.2) and habitat data will be recorded on the appropriate habitat

data collection forms (Appendices C.1, C.2, or C.3) in this book.

LITERATURE CITED

Pulket, M. (editor). 2017. Wadeable multihabitat stream macroinvertebrate data




Pennsylvania.

3-33

3.6 MACROINVERTEBRATE LABORATORY SUBSAMPLING AND

IDENTIFICATION PROTOCOL

3-34

Prepared by:

Dustin Shull






2017

R script provided by:

Zachary Smith

Interstate Commission on the Potomac River Basin

30 West Gude Drive Suite 450

Rockville, Maryland 20850 USA

Edited by:

Mark Brickner






2020

3-35

INTRODUCTION

This laboratory subsampling and identification protocol applies to the freestone riffle-

run, limestone (with exception of subsample target), multihabitat, and semi-wadeable

macroinvertebrate collection protocols. The limestone collection protocol is currently the

only macroinvertebrate collection protocol with a subsample target of 300 ± 30

organisms. All other macroinvertebrate collection protocols described above have a

subsample target of 200 ± 20 organisms.

SUBSAMPLING PROCEDURES

Laboratory Subsampling

Subsampling procedures are adapted from Barbour et al. (1999). Subsampling should

take place in a controlled environment with good lighting, access to water, and plenty of

workspace. Equipment and supplies needed for the benthic sample processing are:

• At least 2 large laboratory pans gridded into 28 squares. The specific size of a

pan is not critical, but the number of grids (28) must be maintained if any basic

density comparisons wish to be made between samples. DEP Central Office staff

use Rubbermaid 2 Gallon White Durex® Container 18" x 12" x 3 1/2", because

the bottom of the pan is 14" x 8", which allows for 28, 4in2 grid cutters to be used

• A standard USGS No. 35 sieve (or 500 µm mesh sieve bucket) to remove fine

materials and residual preservative prior to sub-sampling

• A random number generator or uniform objects (numbered from 1 to 28) for

drawing random numbers

• Forceps (or any tools that can be used to pick organisms and debris)

• Grid cutters that approximate 1/28th of pan surface area.

• A lighted magnifier (10x magnification maximum) may also be used during the

sorting process but is not required. This equipment can make seeing organisms

among debris easier and is good for training, but it does not make

microorganisms (not visible with the naked eye) visible.

• Handheld counter

• Dissecting microscope (primarily used for identifications, but also used to confirm

if organisms are identifiable).

• Storage vials for subsampled organisms

• 70-90% ethanol

• Petri dishes

To begin subsampling, the composited sample is removed from the collection container

and placed in the first gridded pan (pan 1). It is recommended to remove fine materials

3-36

and residual preservative prior to subsampling by rinsing the sample through a sieve (or

sieve bucket) that matches the mesh size (or smaller) of the net gear used to collect the

sample. The sample is then gently distributed evenly throughout pan 1. Water may be

added to the pan before distributing to reduce damage to the organisms.

At this point, there are several ways to achieve the targeted subsample of 200 ± 20 (300

± 30 for limestone stream samples) organisms. It is important to note that this protocol

is a 200 or 300 organism target; thus, collectors should avoid consistently biasing

toward the lower limits of 180 organisms (270 for limestone stream samples) or upper

limits of 220 (330 for limestone stream samples) organisms in the final identifiable

count. Depending on the nature of the sample and the abundance of organisms, one

subsampling procedure may be more efficient than another. The subsampling

procedure is acceptable if it remains random/unbiased and the number of grids in pan 1

and pan 2 can be recorded. Only identifiable organisms (to the taxonomic level

recommended in the identification section below) are counted during subsampling.

Organisms that are not considered identifiable include pupae, larval bodies missing too

many critical structures to render confident identifications, extremely small instars,

empty shells or cases, and terrestrial organisms. If an organism is lying between two

grids it is considered in the grid that contains the head. If the head is not easily seen,

then the organism is considered in the grid containing most of the body. If a large

volume sample necessitates the use of two first pans, material and organisms are

removed from the same grid of both pans. For example, a large amount of debris is

distributed evenly into two first pans. Then, if grid 17 is randomly selected, grid 17 must

be pulled from both pans and sorted. These two grids are then counted as 1 grid from

the first pan.

The following describes two potential ways of conducting randomized subsamples:

Option 1: (good for suspected high abundance samples)

The following option requires three pans (2 gridded pans and 1 viewing pan).

Randomly select 1 grid from the first gridded pan (pan 1) and remove all material and

organisms. Place material and organisms in a viewing pan (pan should be as large as

gridded pans to facilitate quick identification of organisms from debris). If the number of

organisms (through a cursory count) appears to be much greater than 50 (e.g., 75

organisms), then move the material and organisms into the second gridded pan (pan 2).

Randomly select 3 more grids from pan 1 and place all material and organisms evenly

throughout pan 2. This process quickly confirms a high likelihood that there will be more

than 220 (330 for limestone stream samples) organisms in the first 4 grids of pan 1 and

3-37

a second gridded pan will be needed. This process also significantly reduces the

amount of debris a collector must sort through before determining a second pan is

needed. Once material and organisms are evenly distributed throughout pan 2, grids

are randomly picked until the target number of organisms is reached. The total number

of grids for each pan is recorded on the laboratory bench sheet (Appendix A-2) or

annotated in the appropriate section of a mobile data entry application.

Option 2: (good for suspected low abundance samples)

The following only requires 2 pans (1 gridded pan and 1 viewing pan) if the sample has,

in fact, a low abundance of organisms.

Randomly pick 1 grid at a time from the first gridded pan (pan 1) and place all material

and organisms in a viewing pan to be sorted. Once all organisms are removed, the

debris can be discarded. This allows for the viewing pan to be clear of debris for the

next randomly selected grid from pan 1. Continue subsampling 1 grid at a time until the

target number of organisms is reached. With low abundance samples, more than 4

grids from pan 1 will be selected, so there will be no need to use a second gridded pan.

The total number of grids for each pan is recorded on the laboratory bench sheet

(Appendix A-2) or annotated in the appropriate section of a mobile data entry

application.

Once subsampling is completed, all organisms are placed in a clean vial with fresh

70%-80% ethanol. The vial is labeled using indelible ink with date, time, collector,

collectively referred to as a station ID (yyyymmdd-HHMM-Collector), and location and

then stored for later identification. If more than one vial is needed then each vial will also

be labeled with a number (e.g., 1 of 2, 2 of 2). In addition, if identification occurred

immediately after subsampling, then organisms can be placed into a petri dish instead

of a labeled vial.

There will be rare occasions when subsampling results in more than 220 organisms (or

330 organisms for limestone samples) at the end of the subsampling process. If this

occurs, pour the organisms out of the vial and into a clean gridded pan. Evenly

distribute the organisms with fresh ethanol and then randomly select grids to remove

organisms until the target number is achieved. This can be done in a backwards fashion

to save time. For instance, a collector followed the procedures above, but got 244

organisms in 4 grids total. The collector then poured and evenly distributed organisms

into a clean gridded pan. After removing organisms out of 4 randomly selected grids,

the collector had 208 organisms in the gridded pan and 36 organisms set aside. At this

point the collector simply needed to place the organisms in the gridded pan back into

3-38

the vial. The 36 organisms left over were placed back into the original sample jar.

Because the collector subsampled in a backwards fashion, the count of grids in pan 1

remains “4”, but the 4 grids removed during this additional process are subtracted from

28 in a “pan 2” to get 24 grids.

Digital Subsampling (Rarefaction)

Rarefaction Process

Digital subsampling or “rarefaction” is used to reduce sample size of a previously

collected and processed sample. Rarefaction should not be considered a substitute for

the subsampling procedures described above, but this process is acceptable when the

physical sample is unavailable, or an immediate answer is needed. The concept of

rarefaction was developed by Howard Sanders (Sanders 1968), modified by a number

of other scientists (i.e., Hurlbert 1971, Heck et al. 1975, Simberloff 1978). Rarefaction

has since been incorporated into several computer programs. Normally, rarefaction

programs use the original sample numbers and, using a specified subsample amount,

produce the expected number of species and a variance of this expected number of

species. Depending on the program, confidence limits and plots/curves may be

produced. Two important aspects of any rarefaction program are that, 1) the program

can randomly reduce the number of individuals in each taxon, and 2) the program can

randomly reduce sample richness. This ensures that the program functionally replicates

the physical subsampling process.

For conducting rarefaction using spread sheets or designing a rarefaction program, the

following steps should be followed:

1) Calculate how many macroinvertebrates would be in each grid of a 28-grid pan,

assuming they are spread evenly across the pan.

2) Calculate the number of macroinvertebrates to remove based on the desired

subsample (e.g., target for freestone IBI = 200 ± 20 individuals).

3) Divide the number of macroinvertebrates to remove by the number of

macroinvertebrates found per grid.

4) Round this result to the nearest whole number grid.

5) Multiply the number of macroinvertebrates per grid by the number of grids to

subsample.

6) Round this result to the nearest whole number (because entire organisms are

removed, not partial). This is the number of macroinvertebrates that will be

removed from the sample to reach a correct subsample. The remaining

macroinvertebrates will be the correct subsample.

7) Number each individual macroinvertebrate from one to the total number of

individuals in the sample on a spreadsheet.

3-42

made to have other taxonomists assist with a positive identification. Otherwise, the taxa

should be included on the enumeration sheet, separate and distinct from other

identifications within the same taxonomic group. Other orders may be identified to the

lowest level attainable if they are immature and do not exhibit the characteristics

necessary for confident genus level identifications; however, this should be rare if the

proper subsampling process is followed. Identifications with a mixture of higher and

lower taxonomic levels within the same taxonomic group will skew sample scoring and

may result in the sample not meeting quality assurance standards. If identifications with

a mixture of higher and lower taxonomic levels within the same taxonomic group cannot

be avoided, higher level identifications will not be used to calculate metrics. Eliminating

these individuals from the sample avoids artificially inflating richness metrics. In

addition, the sample and the identifications must be forwarded to a certified taxonomist

for further evaluation, and the identified organisms should be preserved and maintained

indefinitely.

Table 3. The taxonomic groups that are identified to a higher taxonomic level than

genus.

Group Identification Level

Midges Family

Snails Family

Clams Family

Mussels Family

Flatworms Phylum (Turbellaria)

Aquatic Earthworms & Tubificid Worms Class (Oligochaeta)

Leeches Class (Hirudinea)

Proboscis worms Phylum (Nemertea)

Roundworms Phylum (Nematoda)

Moss Animalcules Phylum (Bryozoa)

Water mites Subclass (Acari)

All organisms are identified and enumerated on a bench sheet (Appendix A.2) or data

entry application. Organisms are then returned to the labeled vial and stored in a safe

location.

Quality Assurance

At least 10% of all samples identified by a biologist are quality assured by a certified

taxonomist. The most common type of certification is the genus level EPT certification

provided by the Society of Freshwater Science. Identification results between the

biologist and certified taxonomist must be 90% or greater. DEP may request sample

vials for any results entered into internal databases for quality assurance purposes.

3-43

LITERATURE CITED





Heck, K. L., G. Belle, and D. Simberloff. 1975. Explicit calculation of the rarefaction

diversity measurement and the determination of sufficient sample size. Ecology

56:1459–1461.

Hurlbert, S. H. 1971. The nonconcept of species diversity: A critique and alternative

parameters. Ecology 52:577–586.

Sanders, H. L. 1968. Marine benthic diversity: A comparative study. The American

Naturalist 102:243–282.

Simberloff, D. 1978. Use of rarefaction and related methods in ecology. ASTM STP 652

in K. L. Dickson, J. Cairns, and R. J. Livingston (editors). Biological data in water

pollution assessment: Quantitative and statistical analyses. American Society for

Testing and Materials.

3-44

3.7 FISH DATA COLLECTION PROTOCOL

3-45

Prepared by:

Timothy Wertz






2017

Edited by:

Timothy Wertz






2021

3-46

INTRODUCTION

Fish community assessments are important as they represent a biological community

that are captive to water resources during all life stages, have longer life spans, and

have a greater socioeconomic status than macroinvertebrate communities alone.

Standardized semi-quantitative sampling protocols can be applied to all wadeable and

non-wadeable streams. The data generated from standardized protocols are vital for

making ALU assessments and evaluations, developing indices and other assessment

methods, promoting spatial and temporal trend analyses, and providing the foundation

for a robust fish bioassessment program. The Assessment Book (Shull and Whiteash

2021), Chapter 2, includes the Stream Fish Assemblage Assessment Method (Wertz

2021) which requires strict adherence to the following standardized collection protocol.

Other collection protocols may be considered for various assessment purposes only

after rigorous analysis for similarity.

BASIC REQUIREMENTS

Field sampling procedures, species identification and enumeration, and other site tasks

must be applied consistently with the same level of rigor at each sample site. To

maintain consistency, a field crew is directed on site by a crew leader who maintains a

valid scientific collector permit (SCP) as required by PFBC and has met the quality

control requirements of DEP (described below). Accurate species level identification of

each fish collected is essential, as it provides the foundation for subsequent

assessment. A minimum of one crew member must be proficient in the accurate

identification of Pennsylvania fishes and should also be versed in fishes of the

surrounding areas and potential exotics. Electrofishing techniques are the primary

procedures adopted by DEP. Other fish collection procedures are not discussed here.

Electrofishing crews will have a crew leader trained and experienced in the proper

operation of electrofishing gear and appropriate electrofishing safety and strategy

(described below).

REPRESENTATIVE SAMPLING

The objective of this semi-quantitative fish data collection protocol is to acquire a

representative sample of the overall fish community, here referred to as the fish

assemblage. Fish assemblage data is considered representative when it is collected

with a standardized collection protocol that is robust enough to produce repeatable

biological response(s) to environmental and/or anthropogenic conditions. The collection

protocols may or may not provide a complete inventory of all species present at a site

3-47

but should be able to consistently characterize most species present and their relative

abundance.

Sampling Considerations

Sampling effort is measured by both distance and time and accurate measures of both

are considered essential. Accumulated electrofishing time can legitimately vary over the

same distance as dictated by cover, stream conditions, and the number of fish

encountered. While it is understood that some individual fish will not be captured, a

concerted effort by the crew members should be made to capture every fish sighted as

this is fundamental to relative abundance measures. Dipnet design and mesh size will

largely dictate the size range of fishes captured; frame diameter, net depth and handle

length should be of sufficient size to capture all anticipated fishes and size ranges. Net

mesh should be approximately 6mm (1/4in). Since the ability of the netters to see

stunned and immobilized fish is partly dependent on water clarity, sampling is to be

conducted only during periods of “normal” water clarity and flows (e.g., baseflow

conditions). Sites that are turbid should not be sampled if water clarity is under one

meter unless this clarity is considered normal. The sampling period for all semi-

quantitative fish collection protocols is from June 1st through September 30th and this

period may provide a wide variety of stream flow conditions. Periods of high turbidity

and high flows should be avoided due to their negative influence on sampling efficiency.

If high flow conditions occur, sampling must be delayed until flows and water clarity

return to seasonal norms. The crew leader will decide if conditions are acceptable and

direct their crews accordingly. A representative fish assemblage is not only based on

conditions but also on sample reach and habitat selection. A sample reach at a site is

considered most representative when it;

• Contains all the best available habitat (BAH) types (geomorphic units, flow

regimes, fish cover, and substrate material) within a contiguous reach.

• Is similar to that of the local area (i.e., unique habitats should be avoided).

SITE SELECTION

Two semi-quantitative sampling procedures, wadeable and non-wadeable, are

described below. Strict a priori designations of wadeable or non-wadeable have been

avoided due to complex limitations, but see Flotemersch et al. (2006) for a detailed

description of these designations used elsewhere. A stream reach at a site is

considered wadeable or non-wadeable not by size, but in its ability to be efficiently

surveyed during the June 1st -September 30th sampling season utilizing the following

collection protocols (a stream reach may be considered for both protocols throughout

the same period if both protocols can efficiently be implemented). Sites that can have

3-48

both protocols efficiently implemented are referred to as “transitional zones” and are

typically large wadeable sites and small non-wadeable sites (Flotemersch et al. 2006). If

a site is considered within a transitional zone both protocols should be applied within a

short timeframe (days-week), as feasible, to facilitate direct comparisons between

collection protocols. Sites that are too large for efficient coverage using wadeable

techniques (large rivers) are not considered transitional and should only be sampled

using non-wadeable techniques. Duplicates (a second sample reach at a site

comparable to the first, sampled the same day) and replicates (repeating the same site

within the same sampling season) should be performed as often as feasible (preferably

1 out of every 10 sites) for both collection protocols to maintain a calibration dataset for

future assessments.

Final Site Determination

Specific site locations will first be determined by the type of study and its design (e.g.,

cause and effect, protected use assessment) secondly by access feasibility (e.g., public

access, private landowner permission, distance to stream etc.) and lastly by

representative habitat. Desktop reconnaissance as well as a field reconnaissance

should be implemented prior to sampling to determine the appropriate gear and

manpower needed to achieve a representative sample. Further consideration should be

applied to the site selection process to ensure a representative sample of the conditions

that are being measured. Sites located near the mouth, near an instream dam or

impoundment, or near the mouth of a feeding tributary should be avoided unless these

influences are inherent in the study design. General guidance for distances represents

modifications from literature and preliminary data exploration (Gorman 1986, Schaefer

and Kerfoot 2004, Hitt and Angermeier 2008, DEP unpublished data) and are provided

in Table 1. In any case, potentially influencing factors should be noted on the field forms

to facilitate data management and analysis. Applying a detailed reconnaissance to the

site selection process will greatly reduce down-time the day of the sampling event,

ensure repeatability and provide high-quality data.

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Table 1. General guidelines for site selection based on distance from potential

influencing factors.

Influence Category

Distance

(km) Description

In-stream dam and

its impoundment Minor 0.5

- Defined by dam NOT restricting fish

passage under normal to high flow

conditions.

In-stream dam and

its impoundment Major 1.5

-Defined by dam restricting fish passage

under all flow conditions.

Proximity to mouth Minor 0.5 - Defined as target stream within one

stream order of receiving stream.

Proximity to mouth Major 1.5 - Defined as target stream being at least

two stream orders smaller than

receiving stream.

Proximity to tributary Minor 0.5 - Defined as tributaries within three

stream orders of receiving stream.

Proximity to tributary Major 1.5 - Defined as tributaries three stream

orders smaller (adventitious). Should be

avoided within the reach, when feasible.

WADEABLE PROTOCOL

For semi-quantitative fish sampling in wadeable streams, sample reach lengths will be

determined by the width of the stream. The average wetted width is multiplied by 10,

with a minimum reach length of 100 meters and a maximum of 400 meters (Table 2).

Sample reach width is determined by measuring five wetted channel widths spaced

twenty meters apart within the first 100 meters of the proposed sample reach, and

averaged. Once the wetted width has been calculated, it is best to establish the

stopping point before establishing the starting point. Due to the nature of this protocol,

fishes will naturally be “pushed” upstream to avoid the electrical field. The stopping point

should subsequently be a section that provides a natural barrier to escape (e.g., shallow

riffle or cascade) and should be established first. Sample reach length can then be

measured in a downstream direction to determine the starting point. For consistency,

the reach should adhere to the minimum reach lengths but can be lengthened if

necessary, to include additional habitat types. The use of downstream blocknets should

be avoided based on the size range of wadeable streams in Pennsylvania and by the

bias that may be inherent in blocknetting small streams but not large.

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Table 2. Sample reach lengths based on stream width.

Average Stream Width Minimum Reach Length

<10m 100m

10 to 40m 10 X average stream width

>40m Maximum of 400m

Before sampling, the crew leader will brief the electrofishing crew on safety and strategy

(discussed below) and will adjust the electrofishing equipment to the desired settings

(Appendix A.3). When the crew is ready to begin, the crew leader will take note of the

GPS location, will start the clock, and electrofishing will commence in an upstream

direction. The electrofishing crew will be positioned along a transect, adequately

covering the width of the stream, and should maintain this position while proceeding

upstream to avoid gaps in the electrical field which may decrease coverage. The crew

leader must be cognizant of this coverage throughout the survey and adjust as needed.

For best coverage and standardization of effort the rule-of-thumb for determining the

amount of gear needed at a reach is one probe for every five meters of stream width

(Figure 1). For the large wadeable streams this will inherently require a great deal of

planning to ensure the best coverage is achieved (e.g., equipment and manpower). If

gaps are still present along the stream width, the crew can adapt by doing small zigzags

as they proceed upstream for adequate coverage. Similar collection protocols that may

employ transect zigzags across the width of the stream (see, Lazorchak 1998), multiple

pass (see, Pusey et al. 1998, Meador 2003, Shank et al. 2016), or other variant, will not

be considered comparable until a detailed analysis can be performed. If a stream width

is too large for best coverage, nonwadeable techniques should be pursued. Each crew

member with an anode probe should continuously sweep the probe side to side for best

coverage. The number of nets to anode probes must be a minimum of one to one,

additional netters may be used as needed. When the crew reaches the predetermined

stopping point, the crew leader will instruct the crew to stop and take note of the

stoppage time and GPS location. Fish processing will then commence and is described

in detail below.

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extra reach for every 30K sq/km of drainage area over 30K sq/km. For example, if a site

is less than 30K sq/km, RDB and LDB would be sampled, if a site is 30-60K sq/km a

middle reach would be included. Strict definitions are avoided to provide the crew leader

with discretion for best coverage.

Sample reaches are surveyed using one-pass, daytime electrofishing. Reaches are

surveyed in a downstream direction for a longitudinal distance of 500 meters for a

minimum of 30 minutes. Specific sample reaches should be established by desktop and

field reconnaissance prior to sampling. Measures of stream width may be problematic

on large rivers based on rangefinder capabilities and may have to be performed digitally

during the desktop reconnaissance. To maintain strict adherence to the reach length

requirements, starting and stopping locations should be adequately marked with

flagging tape or other distinguishable features in the field prior to sampling. Before

sampling, the crew leader will brief the electrofishing crew on safety and strategy

(discussed below) and will adjust the electrofishing equipment to the desired settings

(Appendix A.3). When the crew is ready to begin, the crew leader will take note of the

GPS location, will start the clock, and electrofishing will commence. A single boat-

mounted electrofishing unit is maneuvered in a zigzag fashion from littoral shoreline

areas toward the thalweg for a distance of 30 meters from the shoreline or to where

stream depth reaches six meters, whichever comes first. If a mid-channel reach of the

river is to be sampled, island habitat must be targeted to satisfy the littoral shoreline

requirement. For non-wadeable sampling, BAH is both representative of the local

conditions, and within a contiguous 500-meter reach. The habitat should be uniformly

navigated and electrofished based on the proportional abundance of the habitat within

the reach. Prolonged electrofishing within a single habitat type is not recommended. If a

small section of the reach cannot be adequately surveyed due to navigational obstacles

or flow restrictions, the area can be avoided and the same distance added to the end of

the reach (electrofishing time must be adjusted accordingly). Best effort should be made

to net all fishes observed and in proportion to their abundance. The strict targeting of

the largest fishes must be avoided.

Each boat crew will have one navigator (usually the crew leader), and a minimum of one

netter for each anode “dropper”. Additional crew members may be utilized to aid in

netting large fishes and monitoring fish stress levels. When the crew reaches the

predetermined stopping point, the crew leader will instruct the crew to stop and take

note of the stoppage time and GPS location. Fish processing will then commence and is

described in detail below.

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FISH HANDLING AND PROCESSING

General Considerations

Since semi-quantitative fish surveys depend on the collection of fish for species

identification, temporary retention of a significant number of fish is required. To minimize

stress and lethality to the greatest extent possible, the following field procedures should

be followed:

• Remove stunned fish from the electrical field as soon as possible.

• Netted fish should be transferred to containment devices as soon as possible.

Containment devices vary depending on the type of survey and situations but

may be a combination of:

o Large buckets or tubs

o Live-wells constructed of framed netting that form a cage that is

submerged in the stream

• Water is replaced regularly in warm weather to maintain adequate dissolved

oxygen levels in the water, reduce waste by-products, and minimize mortality.

• Aeration should be provided to further minimize stress and mortality if necessary.

• Field identifications and releases should commence immediately after collection

• Every feasible effort will be made to minimize holding and handling times.

Special handling procedures may be necessary for species of special concern as

outlined by the SCP requirements. Fish that are not retained for vouchers or other

purposes are released back into the water after they are identified to species and

enumerated. Invasive species will be kept and appropriately disposed of out of the

water, if requested by SCP requirements. Each sample crew must have at least one

person qualified as a taxonomic specialist in field fish identification and must also be

trained in the identification of gross anomalies inherent to the fish health assessment.

FISH HEALTH ASSESSMENT

Incidence and prevalence of gross anomalies are important measures of fish health that

have been used as an indicator of anthropogenic stress (Bauman et al. 2000) and allow

for both spatial and temporal comparison. The measures of deformities, erosions,

lesions, tumors and parasites (DELTP) are completed immediately after sampling. The

intent of the fish health assessment is to record the species, length, and various

anomalies that may be present on each individual fish. This data can then be used to

elucidate any potential issues that may be inherent to a specific species or length-class.

The health assessment is not intended to be diagnostic but should be able to

consistently classify specific gross anomalies by general DELTP type. Because small-

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bodied fishes generally have a shorter life span (or shorter exposure time) the health

assessment is generally intended for fishes >100mm. It is recommended that all fish

>100mm in length be assessed, however, it is understood that large surveys may have

an inordinate amount of fish and may need to be subsampled. Subsampling should be

random (i.e., fish with anomalies should not be targeted), should represent all species

available, and should be of significant sample size relative to their abundance in the

sample. The goal during this process is to record as much information about the fishes

without inflicting undue stress. It is best when fishes that are to be processed for health

are held in live-wells. If the fish appear to be stressed, even after all attempts have been

made to reduce stress, the priority is to release and not collect health assessment data.

When fish are released without health data, only species identification and enumeration

are needed.

To conduct the health assessment the crew leader will assign the following three duties

to the crew: inspector, handler, and recorder. The inspector (generally the crew leader)

should be familiar with common fish diseases, experienced in the identification of fish

and gross anomalies, and have participated in numerous fish health assessments as

either handler or recorder. It is the inspector’s duty to give a species identification,

measure the total length (TL) of the fish (to the nearest 5mm increment), and inspect

the fish for anomalies. TL is measured from the tip of the snout to the end of the slightly

compressed caudal fin. The fish will be inspected along the right and left sides of the

body, head, fins, and gills. Specific anomalies found during this inspection will be voiced

to the recorder. Once the data has been collected, the inspector can release the fish.

The crew leader (or inspector) should determine where fish should be released; fish

should not be released near anticipated sample reaches to avoid immigration and bias

in the next sample. The handler is responsible for monitoring and reducing the stress

level of the fish while in retention and subsampling the fish that are then given to the

inspector. The recorder observes the inspection and records the data voiced by the

inspector. The recorder sets the pace of the health assessment and the inspector

should adjust accordingly. The recorder should be able to anticipate the amount of

space needed to record the data based on the number of fish that are in the sample, as

well as write neatly to avoid error in data management. A quick reference field guide for

anomalies is imbedded in the field data collection form in Appendix A.3.

FISH PRESERVATION

Most captured fish are identified to species in the field and released; however, any

uncertainty about the field identification of individual fish requires their preservation for

future laboratory identification (except for large specimens or species of special concern

which may need to be photographed). Only the fish necessary for vouchering and

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laboratory confirmation will be retained. Fish are fixed for future identification in buffered

10% formalin. The container is labeled, at minimum, with date, time, waterbody, site

location, and geographic coordinates. Species level identification is required and may

be necessary to the subspecies level in certain instances. If a species level identification

is not possible in the field, voucher(s) specimens should be retained, or properly

photographed, for later laboratory analysis. Proper photographs should include a picture

of the left side of the fish body in the anatomical position, a label with site information

and individual identification number, and taxonomically important characters needed for

species level identification. In any case, strict adherence to SCP requirements and

subsequent annual updates must be considered.

The PFBC’s Division of Environmental Services (DES) maintains the general SCP

requirements as well as Qualified Threatened and Endangered Species (TE) Surveyor

requirements. More information on SCP and TE survey requirements can be found at

http://www.fishandboat.com/Resource/EnvironmentalServices/Pages/default.aspx

DES also has guidance for vouchering, and specific species that may be important to

voucher if encountered.

SITE DATA

Prior to leaving the site, the field collection sheet in Appendix A.3 must be completed,

ensuring that all field-based data is recorded. Location information is critical to ensuring

repeatability of a collection, and detailed descriptions are encouraged. Descriptions of

location should include relative position to a stable landmark. For example,

“downstream of bridge” is ambiguous, whereas “250 meters below SR144 bridge

upstream of Crossfork, PA” ensures repeatability. Water quality measures must follow

guidelines as described in Chapter 4.1, In-Situ Field Meter and Transect Data Collection

Protocol (Hoger 2020). Gear and effort descriptions should be as specific as the

equipment allows and are considered essential to normalizing data for effort. Qualitative

habitat measures (Mesohabitat, Velocity/Depth Regimes, Habitat, and Substrate) are

recorded as a percent-occurrence within the sampled reach and should not be applied

to a larger scale. Habitat measures are considered typical except for Percent of Fish in

Habitat. Here, the percent of fish (the number of individuals regardless of species) that

were netted out of each habitat type within the reach is recorded. This measure is

atypical but provides insight into habitat utilization. This measure is not always required

as it can be influenced by reduced visibility or when fish are “pushed” into another

habitat by the electrical field.

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LABORATORY PROCEDURES

Important Note- All chemicals that are integral to the preservation and long-term

storage of fish vouchers should be treated as hazardous. Chemical specific Material

Safety Data Sheets (MSDS) forms should be referenced and be readily accessible at

the laboratory. Containers should be labeled appropriately, and nitrile gloves worn at all

times. General considerations for vouchering specimens come from Walsh and Meador

(1998) and can be referenced for a more detailed analysis.

Sample Preparation

Before fishes can be safely handled in the lab, they must first be transferred from 10%

formalin fixative and washed. Washing includes soaking fish in fresh (tap) water for a

24-hour period per wash, for a minimum of three washes (three days). Larger samples,

with more biomass, may need additional washes. After the last washing period, the

sample can be gradually (over a period of days) introduced to sequential levels of ethyl

alcohol washes from 35% to 50% to 70%. The ideal concentration for long term storage

is a 70% ethyl solution and should be confirmed with a hydrometer after immersion for

24 hours. Once the sample has been properly washed and preserved, laboratory

identifications can be completed.

Identification and Enumeration

Taxonomic identification and enumeration are the two most important factors for quality

lab data. The first step is to obtain the taxonomic identification. Fishes are identified and

segregated into species groups, or taxonomic units. The second step is to lay each

species out and enumerate them, this provides for a second-chance opportunity to spot

any taxonomic errors. It is best to lay species out for enumeration in smaller groups, by

fives or tens, to facilitate a recount. If a second taxonomic expert is available, it is best

to have them confirm the identification and enumeration, which provides a third and final

opportunity to catch any mistakes. Laboratory taxonomic identifications and

enumerations are then added to the original field data sheet to obtain the final taxa list

and enumerations. Nomenclature should follow the most recent update provided by the

American Fisheries Society (Page et al. 2013). Regional taxonomic keys should be

used for identification and range distribution comparisons. Notable identifications of

species outside of their known range should be documented and verified before data

are considered final. Fishes should be identified to the lowest taxonomic level possible,

to include species, subspecies, and hybrids (with comments on parental genus or

species). If identification to this level is not possible (due to size, condition, or other

variable) the use of genus or possibly family level identification may be considered. To

maintain data consistency, fishes <25mm (TL) should not be included in the final

identification and enumeration, but can be included as separate “comments”. For

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species that may have close phylogenetic “sister species” and that can occur together

(sympatric), the documentation of the morphometric or meristic counts used to make the

identification is encouraged.

Sample Retention

Sample retention allows for both short-term and long-term quality assurance and quality

control (QA/QC) of data. In the short-term, samples can be retained for independent

verification or for inclusion into a teaching/reference collection. Sample retention can

also allow for long-term QA/QC of data and can be incorporated into museum records,

usually at a State or Federal institution (Table 3).

Table 3. Contact information for various fish retention organizations. Contact information

subject to change.

Organization Contact Address

Penn State

University Fish

Museum

Dr. Jay R. Stauffer, Jr.

432 Forest Resources

Building

University Park, PA 16802

Pennsylvania Fish

and Boat

Commission

Doug Fischer

PFBC Centre Region Office

595 E. Rolling Ridge Dr.

Bellefonte, PA 16823

Cornell University

Museum of

Vertebrates

[email protected]

159 Sapsucker Woods

Road

Ithaca, NY 14850-1923 USA

The Academy of

Natural Sciences of

Drexel University

http://www.ansp.org/about/staff/contacts

1900 Benjamin Franklin

Parkway, Philadelphia, PA

19103

ORSANCO Jeff Thomas 5735 Kellogg Avenue

Cincinnati, Ohio 45230

National Museum of

Natural History Dr. Jeffrey T. Williams

Div. of Fishes, Museum

Support Center MRC-534

4210 Silver Hill Road

Suitland, MD 20746

Ohio State University Marc Kibbey 123 Derby Hall

Columbus OH, 43210

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To fulfil the requirements of this protocol, a minimum of one “Site Composite Voucher”

is needed for every site, and a second “Audit Voucher” may also be needed as

described below.

• Site Composite Vouchers- A site voucher is for verifying species

spatiotemporal occurrence at a known site. A site voucher is comprised of a

minimum of one individual representing each of the lowest taxonomic units

identified in the sample. At sites with multiple reaches (e.g., nonwadeable) only

one site voucher is needed, but should represent the lowest taxonomic units

combined across all reaches. Not all species will necessarily need to be

vouchered. Larger-bodied fishes, gamefish, or generally common species may

be excluded if desired. These vouchers are labeled using waterproof, chemical-

resistant paper (placed inside the jar) with a minimum of:

o Site- Waterbody and location

o Date- Either Date/Time or a DEP GIS key in the form of

YYYYMMDD-TIME-NAME (Name = Initial of first name and full last

name “JSmith”)

o GPS Location- Latitude, Longitude

o Type- “Site Composite”

o Taxa list- The taxa list should mirror the completed field and lab

compilation and should include the number of specimens

vouchered out of the total number identified (e.g., Moxostoma sp.-

5 of 5, Percina maculata- 1 of 11, Micropterus dolomieu- 0 of 25).

• Audit Vouchers- An audit voucher is for QA/QC of the taxonomic identification

and enumeration. An audit voucher is comprised of all individuals identified and

enumerated at the lab. All fishes identified in the lab are segregated into distinct

containers (or separated within a few containers by a permeable membrane) by

their respective lowest taxonomic unit and are shared with the DEP auditor.

These vouchers are labeled using waterproof, chemical-resistant paper (placed

inside the jar) with a minimum of:

o Site- Waterbody and location

o Date- Either Date/Time or a DEP GIS key in the form of

YYYYMMDD-TIME-NAME (Name = Initial of first name and full last

name “JSmith”)

o GPS Location- Latitude, Longitude

o Type- “Audit Sample”

o Container #- “__ of __”

o Species- Lowest taxonomic unit(s) contained in the jar

(enumeration is not needed for each species, and is described in

detail below).

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QUALITY CONTROL

The QA/QC of data collection is needed to maintain consistent application of these

protocols. This will enhance the precision and accuracy of assessments. The project

coordinator (DEP Quality Assurance Officer), will perform audits on both the field and

laboratory data collection before these data are entered in the DEP database. For field

data collection, a minimum of one audit for each protocol (e.g., wadeable and non-

wadeable) will be conducted for each crew leader. For laboratory data collection, a

minimum of one (but possibly as high as 10% of samples per year) “Audit Sample” will

be required for each taxonomic expert.

Important Note- It is understood that there may be some inherent subjectivity in

defining the “lowest taxonomic unit”, as this is likely to vary between taxonomic experts.

The goal of the laboratory audit is to make these data as consistent as possible, across

the range of expertise. To this end, the project coordinator may require multiple audit

samples before this consistency can be achieved. The auditor may have additional

taxonomic experts perform the laboratory audit to increase consistency. Final

determinations of similarity and consistency within the laboratory audits will be made by

DEP.

Original, signed audit forms will be retained by the auditor and copies will be sent to the

auditee for record. Audit forms for field and laboratory data collection are available upon

request.

ELECTROFISHING SAFETY

DEP recognizes that electrofishing is an inherently hazardous activity for which safety is

a primary concern. The environmental conditions under which these operations are

conducted further increase the risks. Due to these important concerns for electrofishing

safety, all field team members should be familiar with electrofishing techniques, safety

precautions, and equipment manufacturer’s operating procedures. It is the responsibility

of the project coordinator (or crew leader) to verify that crewmembers have a basic

understanding of these principles prior to sampling. The safety of all personnel and the

quality of the data are assured through the adequate education, training, and

experience of all members of the fish collection crew. To this end, the crew leader must

have completed electrofishing training and safety courses, be certified in CPR, and

should have a minimum of five years of electrofishing experience. The US Fish and

Wildlife Service offers educational guidance and certification in electrofishing principles

and safety and is considered the standard training source for electrofishing. Currently,

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US Fish and Wildlife Service offers two courses that are available online. The

Electrofishing Principles and Techniques course is required for crew leaders and the

Electrofishing Safety course should be completed by every crewmember prior to

sampling.

• Principles and Techniques of Electrofishing (Online) - CSP2C01

o https://nctc.fws.gov/courses/CSP/CSP2C01/resources/

• Electrofishing Safety (Online) - CSP2202

o https://training.fws.gov/courses/csp/csp2202/resources/index.html

Electrofishing protocols rely on one of three main types of gear platforms: backpacks

and towboats with hand-held probes for wadeable streams and standard boats with

fixed-boom probes for non-wadeable streams and rivers. Collection gear consists of a

wide variety of net sizes, shapes, and configurations. Each team member must be

insulated from the water and the electrodes; therefore, non-breathable waders are

necessary as they are considered personal protective equipment (PPE), and rubber

(linesman) gloves are recommended. Nonbreathable waders or rubber boots are also

considered PPE and may be worn during boat electrofishing at the discretion of the

crew leader. All waders should have non-slip, rubber soles (no felt soles). Electrode and

dip net handles must be constructed of non-conductive materials (fiberglass or wood).

Additional equipment used during the survey should also be constructed of non-

conductive materials to reduce inadvertent shock potential (e.g., buckets, live-well

frames). Electrofishing units must be equipped with functional safety switches and

represent best available technology (BAT). Field crew members must remain cognizant

of their surroundings and not reach into the water unless the electrodes have been

removed from the water or the electrofishing unit has been disengaged. A good

anacronym to remember for safety is S-H-O-C-K-E-D.

Safety Brief- Crew leaders brief the crew on safety and strategy prior to each collection.

Handle- Be aware of what is touched, stepped on, or ducked under.

Operate- Be aware of probes and the electrical field at all times.

Communicate- Let others know of any potential hazards that may become apparent.

Key Personnel- Know which crew members are trained in case of emergency.

Equipment- Inspect equipment for damage before and after each sampling event.

Debrief- Discuss and critique the survey for improvement.

Each crewmember should also wear polarized glasses to reduce surface glare which

enhances their ability to safely negotiate the stream bottom and to see and net fish.

Important Note- All chemicals that are integral to the preservation and long-term

storage of fish vouchers should be treated as hazardous. Combustible materials used

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for equipment (e.g., gasoline) should also be considered hazardous. Chemical specific

MSDS forms should be referenced and be readily accessible while conducting fish

surveys.

ELECTROFISHING STRATEGY

The efficiency of electrofishing is largely determined by the user’s knowledge of the

equipment. Due to the vast amount of gear types and manufacturers available, it is

imperative that the user is familiar with the gear and the manufacturers

recommendations specific to each unit. Here, only general guidelines are outlined to

standardize capture efficiency. The nature of this protocol inherently targets fishes of all

shapes, sizes, and body forms from the smallest of minnows to the largest catfishes. As

body size and shape varies, so does the transfer of electricity. As size increases

susceptibility to the electrical field also increases. Adjustments of waveform (AC, DC,

pulsed DC), frequency (pulses per second or hertz, Hz) and duty cycle (percent “on” for

a pulsed current) can all have profound effects on the electrofishing efficiency. For

optimal efficiency and consistency waveform, frequency, and duty cycle must be

standardized (Table 4). Environmental conditions, most notably specific conductance,

may require a deviation from the standard waveform from PDC to AC. Using AC is only

advised when specific conductance is very low, and PDC is not able to produce the

desired response.

Table 4. Standardized electrofishing settings

Setting Category Required Setting

Waveform Pulsed Direct Current (PDC)

Frequency 60 Hz

Duty Cycle 25%

The desired response from fish to the electrical field is immobilization with minimal

injury. By standardizing waveform, frequency and duty cycle, the output wattage (amps

x volts) is the remaining adjustment needed to achieve the desired response. Adjusting

the wattage is typically done by adjusting voltage (refer to the manufacturers

specifications for wattage adjustments, as many of these adjustments vary across

units). Testing for the desired response should be done prior to beginning the survey,

outside of the survey area. To this end, output wattage is set to minimum and gradually

increased until the desired response from fish is observed. It is best to use small-bodied

fish for the observed response, as they are generally less susceptible to the electrical

field. A quick reference guide to electrofishing adjustment is in Appendix A-3.

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BIOSECURITY – AQUATIC INVASIVE SPECIES

The spread of aquatic invasive species (AIS) is always a potential concern when

conducting stream surveys. Biosecurity measures should be considered and

implemented for all fish collections. AIS covers a wide range of taxa that include but are

not limited to fish, reptiles, mussels, invertebrates, aquatic plants, algae, pathogens,

and parasites. While many of the streams within Pennsylvania have, or have had

documentation of AIS, it is best to consider all streams as potential candidates for AIS

transfer. To this end, sampling design should be adjusted accordingly. Whenever

possible, site selection priority should be placed on sites within the same watershed and

ordered upstream to downstream. All gear that may come in contact with the water

should be decontaminated prior to leaving the site if they are to be used at another site

within the same day, this includes but is not limited to boats, trailers, live-wells, buckets,

nets, and waders. Various decontamination products are commercially available and

include but are not limited to chlorine solutions or peroxygen solutions. Whenever gear

must be used in multiple watersheds adequate time must be allotted for additional

decontamination practices that include but are not limited to high-temperature pressure

washing, freezing, or drying. Additional information regarding AIS can be found in the

PFBC’s Biosecurity Protocol (available upon request).

LITERATURE CITED

Baumann, P., V. Cairns, B. Kurey, L. Lambert, I. Smith, and R. Thoma. 2000. Lake Erie

lakewide management plan (LaMP). Technical Report Series. LaMP.

Flotemersch, J. E., J. B. Stribling, and M. J. Paul. 2006. Concepts and approaches for

the bioassessment of non-wadeable streams and rivers. U.S. Environmental

Protection Agency, Office of Research and Development, Washington, DC.

Gorman, O. T. 1986. Assemblage organization of stream fishes: The effect of rivers on

adventitious streams. The American Naturalist 128(4):611–616.

Hoger, M. S. 2020. In-situ field meter and transect data collection protocol. Chapter 4.1,




Hitt, N. P., and P. L. Angermeier. 2008. Evidence for fish dispersal from spatial analysis

of stream network topology. Journal of the North American Benthological Society

27(2):304–320.

Lazorchak, J. M., D. J. Klemm, and D. V. Peck (editors). 1998. Environmental

monitoring and assessment program surface waters: Field operations and

methods for measuring the ecological condition of wadeable streams.

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EPA/620/R-94/004F. U.S. Environmental Protection Agency, Office of Research

and Development, Research Triangle Park, North Carolina.

Meador, M.R., McIntyre, J.P. and Pollock, K.H., 2003. Assessing the efficacy of single-

pass backpack electrofishing to characterize fish community structure.

Transactions of the American Fisheries Society 132(1):39-46.

Page, L. M., H. Espinosa-Pérez, L. T. Findley, C. R. Gilbert, R. N. Lea, N. E. Mandrak,

R. L. Mayden, and J. S. Nelson. 2013. Common and scientific names of fishes

from the United States, Canada, and Mexico. Page 243. American Fisheries

Society, Bethesda, Maryland.

Pusey, B. J., M. J. Kennard, J. M. Arthur, and A. H. Arthington. 1998. Quantitative

sampling of stream fish assemblages: Single‐vs multiple‐pass electrofishing.

Australian Journal of Ecology 23(4):365–374.

Schaefer, J. F., and J. R. Kerfoot. 2004. Fish assemblage dynamics in an adventitious

stream: A landscape perspective. The American Midland Naturalist 151(1):134–

145.

Shank, M. K., A. M. Henning, and A. S. Leakey. 2016. Examination of a single-unit,

multiple-pass electrofishing protocol to reliably estimate fish assemblage

composition in wadeable streams of the Mid-Atlantic region of the USA. North

American Journal of Fisheries Management 36(3):497–505.

Walsh, S. J., and M. R. Meador. 1998. Guidelines for quality assurance and quality

control of fish taxonomic data collected as part of the National Water-Quality

Assessment Program. US Department of the Interior, US Geological Survey.

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3.8 FISH TISSUE DATA COLLECTION PROTOCOL

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Prepared by:

Timothy Wertz and Josh Lookenbill






2017

Edited by:

Timothy Wertz






2021

3-66

INTRODUCTION

Routine monitoring for contaminants in fish tissue was initiated in 1979 as part of a

nationwide network of stations encouraged by USEPA. In 1983, Pennsylvania decided

the fish tissue sampling program should be designed to support public health protection.

The USEPA outline for the nationwide program included a list of parameters for analysis

in fish, including PCBs, pesticides and selected heavy metals. This parameter list is

used in Pennsylvania’s fish tissue sampling program. The importance of the fish tissue

sampling and advisory issuance program was fully recognized in May 1986 with the

signing of an interagency agreement between the Department of Environmental

Resources (now DEP), the Pennsylvania Department of Health (DOH), and the PFBC.

This agreement was developed because the agencies desired to pursue a systematic

approach for the detection and evaluation of fish tissue contamination and to develop

procedures for informing the public that may consume such fish of possible adverse

health impacts. The agreement listed the responsibilities of each agency and provided

for the “timely joint issuance of a health advisory” when fish tissue contamination

constituted a health risk. The first joint advisory was issued in June 1986 and included

several waters throughout Pennsylvania. Fish tissue monitoring continues to this day for

303(d) impairment listings and the protection of human health through tiered meal

advisories.

SITE SELECTION

It is important to understand fish tissue evaluations when making site selection as this is

an important component of fish tissue monitoring. To facilitate this understanding, the

term site is a monitoring term for a general area at a local scale and delineation is the

area within the site that’s being evaluated. Herein, a sample is collected at a site that

will be within a previously assessed delineation, unless of course, it is the first sample.

This relationship is important to understand because site selection is largely based on

knowledge of previous assessments and their delineations as established in the

Assessment Book (Shull and Whiteash 2021), Chapter 2, Biological Assessment

Methods. The priority list for determining fish tissue sampling sites is provided in Table

1.

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Table 1. Site selection priority descriptions.

PRIORITY DESCRIPTION

1

Previously unassessed waterbody, specifically those with questionable water

quality and/or elevated angling pressure

2

Previously assessed delineation, needs a verification (resample) to change a

current advisory

3 Rotations, WQN stations = 5YR, Large River Stations = 2YR

4 Follow-ups, existing advisories based on historic data collections

Once a site has been prioritized, a sampling location must be identified. Sampling

locations should be determined by access feasibility and must also be representative of

the site. For example, if a tributary that flows into a large river has been prioritized as a

site, and the current assessment delineation is the entire sub-basin the sample should

not be collected at the mouth where riverine fish may frequently reside. In this case,

samples should be collected from angler-use areas further upstream that are more

representative of the site.

SAMPLE COLLECTION

Once the site and representative sample location are determined, the next step is to

determine what gear is needed survey the area. The most common protocol is by

electrofishing. Electrofishing gear should be suited to the size of the waterbody at the

site and should be capable of efficient capture. While electrofishing remains the most

common, seines, gill nets, trotlines, and angling may be more suitable in certain

situations. Gear and nets should be kept clean and in good working condition to avoid

dirt, oil, and grease from contaminating the sample.

Species are targeted by their recreational and/or consumptive importance. Generally,

the most common game fishes, that are of legal harvest size, are targeted as they are

the most likely to be consumed. In trout streams, samples should be wild fish or hold-

overs greater than, or equal to, seven inches. The targeted size of the fish within a

sample is an important component to the assessment of the data. Fish tend to

bioaccumulate contaminants so older, large-bodied fish will typically have the highest

contamination level. With only a few exceptions, a sample is a composite of fillets from

three to five individual fish representing the same species, that are similar in size. The

75% rule is described on the back of the fish tissue field data sheet (Appendix A.4), and

states that the smallest fish within a sample should be 75% the length of the largest fish

within the sample. Once enough fish are collected at a site to satisfy the targeted

priorities, sample preparation can begin.

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FIELD SAMPLE PREPARATION

Individual fish within a sample are measured for length to the nearest tenth of an inch

and weighed to the nearest ounce. A health inspection is visually conducted for

DELTP’s using the anomaly reference guide located on the back of the field data sheet.

If multiple samples are collected at the same site, the samples should be processed

separately and on clean restaurant-grade aluminum foil (dull side always contacting the

sample). Latex or nitrile gloves are recommended during sample preparation and

should be changed between samples. Before filet preparation, the filet knife is washed

and rinsed with purified hexane labeled as suitable for pesticide residue analysis. Filet

preparation will vary slightly depending on the species, to mimic the most common

preparation techniques for consumption by anglers. For American Eels, five one-inch

cross sections are removed from a skinned and eviscerated individual. For catfishes,

skinless filets are removed from each individual. For typical scaled fishes, the scales are

removed and the skin remains on the filet. Two filets are usually taken from all species;

however, for eels, the sample then contains six to ten filets. If the sample is comprised

of large-bodied fishes (>12in), only one filet from each fish should be included. Once a

sample is filleted it is wrapped in aluminum foil and taped. To avoid processing errors,

the sample needs to be labeled with a minimum of Station Name, Waterbody, Location,

Date and Time, Species, Collector Number and Sequence Number information. The

wrapped and labeled sample can then be placed in a Ziploc® bag and kept on ice in a

cooler until they can be transported to a deep freezer. It is helpful to double check field

sheets for completeness and consistency with the labeled sample before leaving the

sampling site.

DATA MANAGEMENT

Field data should be entered into the SIS database as soon as possible after the

sample has been collected. General SIS data entry procedures are described in

Chapter 4.6, Sample Information System (SIS) Data Entry Protocol (Shull 2017). Fish

tissue data entry is similar to most SIS data entry, with only a few unique differences

described in Figures 1-3.

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Figure 1. Important SIS data entry blocks for Project/Facility data.

Figure 2. Important SIS data entry blocks and fish tissue button (red arrow), for

Sampling Conditions data.

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Figure 3. Important SIS data entry blocks for Fish Tissue data.

All fish tissue data are tied to the project FISH in SIS to maintain a segregated dataset

for monitoring and assessment purposes. The sampling locations under the project

FISH are either WQN stations or WQF stations. If a new location has been sampled, the

DEP fish tissue coordinator should be advised and can add the new location

information.

SHIPPING SAMPLES

Before samples can be delivered to the BOL for analysis, a laboratory sample

submission sheet (Submission Sheet) must be completed for each sample with the

collection information and SAC needed. Make a note that samples are “Fish Tissue

Samples” in the additional information section of the laboratory sample submission

sheet. A list of common fish tissue SACs are listed in Tables 2-4. SACs should be

chosen based on previous results from the same species (of the same length group if

there is a size range description) from a delineated assessment. If the sample is the first

sample at a location, or if there is reason to believe the fish are subject to

contamination, all reasonable SACs should be requested.

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Table 2. SAC 059 (Metals)

Test Code Test Description

71930 Mercury in fish, wet weight****

71936 Lead in fish, wet weight

71937 Copper in fish, wet weight

71939 Chromium in fish, wet weight

71940 Cadmium in fish, wet weight

71945 Selenium in fish, wet weight

71946 Ba in fish, wet weight

71947 SR in Fish, Wet weight

99014G Fish Preparation and Grinding

Table 3. SAC PESTF (Pesticides in Fish Tissue)

CAS Number Analyte (Test) Description

1024573 Heptachlor Epoxide

2385855 Mirex

26880488 Oxychlordane

309002 Aldrin

319846 Alpha-BHC

3424826 O,P-Dde

3734483 Chlordene

39765805 Trans-Nonachlor

50293 4,4'-Ddt

5103719 Alpha-Chlordane

5103731 Cis-Nonachlor

5103742 Gamma-Chlordane

53190 O,P-Ddd

58899 gamma-BHC (Lindane)

60571 Dieldrin

72208 Endrin

72435 Methoxychlor

72548 4,4'-Ddd

72559 4,4'-DDE

76448 Heptachlor

789026 O,P-Ddt

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Table 4. SAC PCBF (PCBs in Fish Tissue)

CAS Number Analyte (Test) Description

Lipids

EXTRACTED DATE

11096825 Arochlor 1260

11097691 Arochlor 1254

11104282 Arochlor 1221

11141165 Arochlor 1232

12672296 Arochlor 1248

53469219 Arochlor 1242

Before shipping the sample to the lab for analysis, the lab sample submission sheet as

well as the field data sheets should be scanned and forwarded to the DEP fish tissue

data coordinator and the BOL fish tissue data representative. The estimated

shipment/receipt date should be described to ensure lab representatives will be

available to receive the frozen samples upon arrival. Once a shipment/receipt date has

been confirmed the samples are ready to send. Completely frozen samples that are of

considerable mass can be placed in a cooler without additional ice and couriered to the

BOL. Small and medium samples should always be shipped on dry ice. The completed

sample submission sheet should be placed in a separate Ziploc® bag and placed inside

the cooler with the sample. To ensure adequate time for lab preparation, analysis and

quality control of the samples, all samples should be shipped to the lab by November 1st

of the collection-year.

LAB SAMPLE PREPARATION

Once the sample has been received and inventoried at the BOL, the samples will need

to be prepared for analysis. The processing equipment is washed and then rinsed with

purified hexane labeled as suitable for pesticide residue analysis. Clean, restaurant-

grade aluminum foil (dull side contacting the sample) should be placed on all surfaces

the sample contacts. Each sample is ground in a meat grinder until all fillets are

considered homogenous (large samples may need to be ground multiple times). The

sample is then subsampled into four 30-gram aliquots that will be used for the analysis

of each SAC, with a backup for reanalysis or archival as desired. The aliquots are

wrapped in foil, pressed to form a patty, labeled with sample number, and refrozen for

analysis. Equipment cleaning, hexane rinses, and foil changes must be completed

before the next sample can be processed. All samples should be processed for lab

analysis by December 31st of the collection-year.

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LITERATURE CITED

Shull, D. R. 2017. Sample information system (SIS) data entry protocol. Chapter 4.6,






Pennsylvania.

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3.9 MUSSEL DATA COLLECTION PROTOCOL

3-75

Prepared by:

Richard Spear, Daniel Counahan, and Dustin Shull


Southwest Regional Office


Clean Water Program

400 Waterfront Street

Pittsburgh, PA 15222

2017

3-76

INTRODUCTION

Larger rivers and streams are highly complex ecosystems with a great deal of

interdependence between biological communities. As a result, biological components of

this assessment must include a diversity of sampling including, but not limited to fish,

benthic macroinvertebrates, and mollusk sampling. Current mussel data collection

protocols are not suited for a rapid and cost-effective means of assessing a

representative portion of the mussel community. Therefore, DEP developed a mussel

sampling protocol that incorporates random sampling to allow for a semi-quantitative

assessment of mussel communities. The DEP freshwater mussel data collection

protocol enables sampling in reaches where fishes and benthic macroinvertebrates

were sampled to provide comprehensive sampling of aquatic biological communities.

This protocol can also be described as a cursory evaluation of the mussel community so

that future/more intensive surveys can be conducted.

The DEP freshwater mussel data collection protocol is designed to enumerate several

aspects of a mussel community. All live mussels are identified to species and

individuals are counted to obtain a representative description of the community in terms

of relative abundance and species richness. Size measurements such as length, width

and depth can be used to approximate age classes of each species. Additionally, this

survey can be used as a rapid and cost-effective way of searching for rare, threatened,

endangered, and invasive mussels. Protocols such as the ones described by Smith

(2006) and Clayton et al. (2013) are also recommended for this purpose.

This protocol is composed of 12 five-minute dives along linear transects that are

randomly spaced from both the downstream sampling point and the shoreline. The

objective of this survey protocol is to build Pennsylvania’s capacity to assess conditions

of large rivers by providing a foundation for the development of large river assessment

protocols for non-wadeable streams. These large river protocols will provide the

capability to conduct bioassessments and monitor environmental conditions with a

scientifically defensible protocol applicable to Pennsylvania’s rivers and streams.

SAFETY PROCEDURES

DEP recognizes the need to structure and oversee this activity due to the inherent

hazards involved in underwater diving. Due to these important concerns for diving

safety, all dive team members must be trained in mussel sampling techniques, safety

precautions, and equipment manufacturers’ operating procedures. In addition, all

scientific divers must have completed, at minimum, an approved SCUBA Certification

Course, and have adequate training to conduct mussel surveys. It is the responsibility of

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the Unit Diving Officer (UDO) to verify that the team members have the appropriate

training. Specific team member duties should be reviewed during each sampling event.

The safety of all personnel and the quality of the data are assured through the adequate

education, training, and experience of all members of the scientific diving team. At least

one UDO must be involved in each sampling event.

The safety standards which will apply and will be followed for all DEP operations are

derived from USEPA's Diving Safety Manual Revision 1.2, (DSM, USEPA 2010b).

Safety Plan

This safety plan establishes general guidelines and procedures for safe and efficient

scientific diving. The intent is to provide written guidance for quick reference in the field

by the Unit Dive Officer (UDO) and scientific divers and is based on the DEP’s Policy for

Scientific Diving Personnel and Diving Operations (DEP 2012). This plan is comparable

to and was derived from USEPA's DSM, and specifically, Appendix A of the DSM,

USEPA Diving Safety Rules (USEPA 2010a). A copy of this safety plan is required to be

available in the field for all diving operations.

Dive Operations and Rules

1. A Dive Plan for each proposed diving operation will be approved by the UDO.

2. Diving with air will be conducted in accordance with the U.S. Navy No-

Decompression Limits and Repetitive Group Designation Table for No-

Decompression Air Dives, located in Appendix H-10 of the DSM (USEPA 2010b).

Diving with Nitrox will be conducted in accordance with the National Oceanic

and Atmospheric Administration (NOAA) Nitrox Tables for No-Decompression

Limits and Repetitive Group Designation Table for No-Decompression Dives

(NOAA 2016).

3. During "live-boat" diving, the tender or UDO will maintain continuous visual

contact with the divers, or their bubbles after they descend, and keep the boat

operator informed of their position. The boat should always be positioned to

render immediate assistance to the divers.

4. The dive plan should require that the deepest dive be scheduled first and be

followed by shallower dives. If this is not possible, the divers must comply with

rule Nos. 5 and 6, below.

5. Generally, all dives should be terminated 5 minutes before the no-decompression

limit.

6. During repetitive diving at water depths greater than 50 feet, on ascent the diver

should stop at approximately 20 feet for 2 minutes for a safety decompression

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stop on the second and succeeding dives. If a safety decompression stop is

taken, the time spent at the stop must be added to the bottom time for that dive.

7. While diving, divers will remain in contact (visual, auditory, tactile, or through

communications from the tender) with at least one other diver. All divers must

surface if contact or communication is lost. No one may dive unattended.

8. A diver should start their ascent from the bottom when their tank pressure

decreases to 500 psi. A diver must begin an ascent with at least 500 psi in

his/her cylinder if they must make a safety stop at 20 feet or if required by

environmental conditions.

9. When diving in extremely limited visibility and overhead environments such as

piers or other situations presenting entrapment or entanglement hazards, the rule

of thirds should be followed: one third of the tank pressure may be used to get to

the dive objective, the second third may be used to complete the work

assignment and return to the surface, and the final third of the tank pressure is

for a safety reserve. This rule does not supersede the 500-psi rule (Rule No. 8).

10. When diving in areas with possibilities of entrapment or entanglement such as

described above in Rule No. 9, an extra open-circuit SCUBA regulator will be at

the dive site and attached to a standby full SCUBA cylinder complete with

backpack.

11. During dives always ascend at a rate of 30 fpm (minor variations in ascent rate

between 20 and 40 fpm are acceptable).

12. No dive will go beyond 130 feet of depth.

13. Diving operations must be conducted in accordance with all appropriate DEP

policies and standards (e.g., for Nitrox diving or flying after diving).

14. In an emergency, the UDO in charge of the diving operation may have to make

field decisions that deviate from the requirements of this safety plan to prevent or

minimize a situation which will likely cause death or serious physical harm.

15. Oxygen will be provided on-scene and on boats participating in dive operation.

Responsibilities of Dive Personnel

UDO:

1. Preparing for the diving operation including checking equipment such as the

pressure in the O2 cylinders.

2. Ensuring that all divers are physically fit to dive.

3. Assisting the Tender and is completely in-charge of the diving operation.

4. Ensuring that all diving operations are conducted safely in accordance with

USEPA's DSM and standards (USEPA 2010b).

5. Promptly prepares a Dive Report upon completion of the diving activities.

6. Giving the pre-dive Briefing which will include:

a. Designating dive buddy pairs including the alternate UDO.

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b. Providing a brief description of the dive site.

c. Discussing the objectives of the diving operation.

d. Reviewing the operation of equipment to be used.

e. Identifying any potential pollution sources.

f. Discussing environmental and any hazardous conditions.

g. Monitoring divers for signs/symptoms of "bubble trouble."

h. Reviewing emergency and evacuation procedures.

7. For the emergency and evacuation procedures, the following will be identified and

discussed in as much detail as is appropriate for conditions at the dive site:

a. Establish evacuation routes and means of transportation.

b. Review methods of communication.

c. Review CPR and the use of medical O2, if necessary.

8. In the Event of a Diving Accident the following will be conducted:

a. Maintain heart and breathing functions.

b. Do NOT remove O2 from patient unless necessary.

c. Reconstruct dive profiles while in route to the chamber.

d. Ensure dive partner accompanies victim or goes to chamber ASAP.

e. Retain all dive gear for examination.

Scientific Divers:

1. Dive only if they are physically and mentally fit and properly trained for the task to

be performed.

2. Keep their diving equipment in safe operating condition.

3. Wear a compass and depth and tank pressure gauges and, when necessary, a

dive watch or bottom timing device.

4. Refuse to dive if diving conditions are unsafe or unfavorable, or if the diving

operation breaches the dictates of USEPA's safety policies or standards (USEPA

2010a, USEPA 2010b).

Standby Divers:

1. Be fully equipped and ready to give immediate assistance at the dive site.

2. Receive the same briefing and instructions as the working divers.

3. Monitor the progress of the diving operations.

Tenders:

1. Assist divers and track their location in the water.

2. Record each diver's bottom time, tank pressures, and maximum depth.

3. Alert the divers, when necessary, on the status of their bottom time via the Diver

Recall Unit.

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4. Advise other vessels of diving operations and warn off boat traffic which may

pose a hazard to the divers.

5. Perform no other concurrent function which interferes with these duties.

Number of Personnel Per Dive Team

Except under emergency conditions, the minimum number of personnel required per

dive team will be as follows (Table 1).

Table 1. Dive team members required.

Water Depth/Situation

Number

of

Divers

Number of

Standby Divers

Number of

Tenders

Total

Individuals

Under 15ft/diver visible at

all times 1 11/ 1 3

Between 15 and

60ft/without unusual

conditions3/

2 - 12/ 3

Between 15 and 60ft/with

unusual conditions 2 11/ 1 4

Between 60 and 130ft/all

conditions 2 11/ 1 4

1/ The Standby Diver will be the UDO or Alternate UDO, if the UDO is diving. 2/ The Tender will also be the UDO or Alternate UDO, if the UDO is diving. 3/ Unusual conditions as determined by the UDO considering weather, water

currents, visibility in the water, potential entanglements, or any other factor that may

compromise the safety of the diving operations.

Decompression Diving

No decompression diving will be conducted

Dive Buddy

No one may dive unattended. Divers must have visible, auditory, or tactile,

communication with at least one dive buddy or a tender at all times.

Dive Computers

As specified in the Dive Plan, determined by the UDO, and in accordance with the DSM,

dive computers may be used to control the dive profile.

Dive Equipment

All dive equipment will conform to USEPA regulations.

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Flying after Diving: wait a minimum surface interval of 12 hours prior to flying after diving.

When making daily, multiple dives for several days or making a dive requiring an

emergency decompression stop, extend the surface interval beyond 12 hours. Whenever

possible wait 24 hours before flying.

Record Keeping

All diving operations will be logged in accordance with the USEPA diving directives in

Section 3.4 of the DSM (USEPA 2010b) and copies of individual divers’ logs will be

provided to the UDO on the first working day and no more than five working days after

the operations are complete. A complete dive report will be provided by the Divemaster

in charge to the UDO within 10 Working days of completion of the operation.

DATA COLLECTION

Survey Crew and Equipment

A survey crew (or dive team) consists of at least three personnel; the UDO, diver, and

stand-by diver. However, more efficient crews should have one to two dive tenders. The

number of crew members required will depend on the depth and type of diving at the

site. For example, some sites may be surveyed by snorkel and mask, requiring fewer

crew members, whereas diving to depths greater than 30 feet or in low visibility would

require more personnel to safely conduct the survey. The types of dives and the number

of members required are described in greater detail in the safety plan (above).

Equipment needs will greatly depend on weather conditions and sampling environment.

Most mussel surveys will be conducted from a boat. The boat needs to be large enough

to accommodate 3 to 5 crew members and approximately 200 lbs. of equipment. In

addition to the standard boat safety equipment, a dive flag should be added to the

inventory. Weather conditions and water conditions will dictate what type of diving

equipment will be necessary to conduct the survey. Survey sites are identified using a

Global Positioning System (GPS) receiver and laser rangefinder. It can be more efficient

to mark each location with a weighted buoy before dives are started. Equipment needed

during dives and mussel processing includes numbered mesh bags, bucket, camera,

calipers, and scale. A checklist of required and suggested equipment is provided below:

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Dive Equipment

Masks

Snorkels

Wetsuits

Gloves

Boots

Hoods (If Applicable)

Fins (If Applicable)

Buoyance Control Devices (BCDs)

Regulators, Octos And Hoses

Air Tanks

Dive Weights

Communication Gear

Dive Flags

Dive Knives

Dive Lights

First Aid Kit

Emergency Oxygen Kit

Other: __________

Field Meters & Related Supplies

Dissolved Oxygen Meter

Replacement Membrane Kits

Do Probe Solution

Zero % Calibrating Solution (If

Applicable)

Ph Meter

Buffers (Ph 4, 7, 10)

KCl Probe Solution

Conductivity Meter

Calibrating Solution (If

Applicable)

Thermometer

Meter Field Manuals (If Applicable)

Secchi Disk

Other: __________________

Mussel Sampling Equipment

GPS

Laser Rangefinder

Numbered Mesh Bags

Bucket

Calipers (Mm)

Scale (Grams)

Camera

Other: __________________

Forms

Field Survey Sheets

Mussel Data Sheets

Other: __________________

Misc.

Maps

Markers, Pens, & Pencils

Calculator

Insect Repellent

Sunscreen

Screwdriver/Tools

Batteries (D-Cell, Other: ________)

Other: _______________

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Survey Location and Site Selection

Survey locations are typically situated at or near large river fish and macroinvertebrate

sample sites and are along the left bank, right bank, or island shore. At each survey

location, dives are conducted at 12 randomly selected sites in a 500m by 50m reach

(Figure 1). The 12 sites will be selected to proportionally reflect shallow and deep-water

habitats. This is supported by previous work by DEP showing distinct shallow or deep

mussel communities. A minimum of 4 shallow water sites will be selected at each

sample site to ensure adequate coverage of this habitat type. For some survey

locations, mid-channel sites may be limited by commercial or recreational traffic that

could pose unsafe diving conditions. Any intake structures or other potential hazards will

be avoided by an approximate 50m buffer. No transects will be performed between any

lock chamber mooring points and the lock and dam structure. If a survey is located near

a US Army Corp of Engineer lock and dam, then the US Army Corp of Engineer must

be notified prior to the survey date. When on site, the lockmaster must be notified via

marine radio that the dive team is ready to commence operations.

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Mussel Data Collection Procedure

A crew member will use a GPS to identify the downstream point of the survey location.

This site is the “X” site in the field survey sheet (Appendix A.5). A range finder will be

used to determine the location of each of the 12 sample locations by measuring the

distance from the shore and the distance from the “X” site. Once at a site, a Latitude

and Longitude is taken using a GPS, and the stand-by diver will deploy the anchor.

In deeper diving conditions or in low visibility, the diver and stand-by diver will dive

together and at least one of those individuals will have communications with the surface.

In shallow diving conditions or if the diver is visible from the surface the stand-by diver

may stay on the surface, but will maintain auditory or tactile contact with the diver. In both

shallow and deeper dives, the diver will record an upstream compass heading while

onboard for underwater orientation. The diver will also be equipped with a 250-foot dive

reel that is marked in 1-meter increments. The UDO will perform a safety check using

communication gear. The diver will descend along the anchor line to the bottom of the

river with the stand-by diver and clip the dive reel to the anchor.

At the beginning of the survey the diver may collect a bucket excavation sample to check

for the presence of mussels in deeper sediment. The diver will communicate to the stand-

by diver that the time interval should begin. The stand-by diver will keep time. For five

minutes, the diver will travel upstream searching (visual and tactile) a one-meter wide

area and collect live mussels and dead mussel shells in the search area. The diver will

search under any large rocks that can easily be turned. The diver will also sweep across

the top of the fine substrate to feel for mussels which may be partially buried. The

sampling protocol recommends a surface search only; subsurface searching will not be

conducted. At the end of five minutes, the stand-by diver will call to the diver via

communication equipment or other diver recall system to conclude the search. All shell

material, live or dead, will be placed in a numbered bag. On a wrist dive slate or verbally,

the diver will record or communicate the substrate composition percentages for fines,

sand, gravel, cobble, boulder, and bedrock. Invasive mollusks (Zebra mussels and

Corbicula) will be scored: 0 = no invasive mollusks; 1 = 1 individual; 2 = multiple

individuals; 3 = small clumps of individuals; 4 = greater than 50% coverage; 5 = complete

coverage of available substrate. The average depth of the searched area will be recorded.

While returning to the boat, the diver will record or communicate the total distance traveled

in meters by counting the number of marked increments on the dive reel. Distance/area

searched will vary based on visibility, mussel density and habitat.

A crew member will use a cleaned and calibrated water quality field meter to record

temperature, pH, dissolved oxygen, percent dissolved oxygen, turbidity, and specific

conductivity according to Chapter 4.1, In-Situ Field Meter and Transect Data Collection

Protocol (Hoger 2020). A Secchi disk is used to measure the depth of the photic zone.

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Water chemistry measurements will be recorded at one of the 12 sites. All live mussels

are identified to species level on-site. However, some representatives may be taken back

to the laboratory for verification if a photograph is insufficient (State and/or Federal

permits required). Live mussels are then weighed (in grams), measured (length, width,

and depth in millimeters), photographed and returned to the river. Measurements of

length, width and depth are described in Figure 2. When returning live mussels, care

should be taken to orient the mussel posterior side up. All mussel data are recorded on

mussel data sheets provided in Appendix A.5. Mussel shells will be identified if both

valves are present and noted as “dead” on the data sheet. Mussels and shells found near

the sampling sites can be identified, measured if live, and recorded on mussel data

sheets, noted as “incidental.”

A. B.

C.

Figure 2. A. Measurement of length is the maximum distanced between the anterior

and posterior margins. B. Measurement of width is the maximum distance between

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dorsal and ventral margins. C. Measurement of depth is the maximum distance between

left and right valves (shell halves).

LITERATURE CITED

Clayton, J., B. Douglas, P. Morrison, and R. F. Villella. 2013. West Virginia mussel

survey protocols. US Fish and Wildlife Service, US Department of the Interior,

Washington, DC.

DEP. 2012. Policy for Scientific Diving Personnel and Diving Operations. Pennsylvania






NOAA. 2016. NOAA No-decompression tables for Nitrox Dives. NDP 07 -2015. U.S.

Department of Commerce, National Oceanic and Atmospheric Administration,

Office of Marine and Aviation Operations, Silver Spring, MD.

Smith, D. R. 2006. Survey design for detecting rare freshwater mussels. Journal of the

North American Benthological Society 25(3):701–711.

USEPA. 2010a. Appendix A EPA diving safety rules. Diving safety manual (Revision

1.2). U.S. Environmental Protection Agency, Office of Administration and

Resources Management, Safety, Health and Environmental Management

Division, Washington, DC.

USEPA. 2010b. Diving safety manual (Revision 1.2). U.S. Environmental Protection

Agency, Office of Administration and Resources Management, Safety, Health

and Environmental Management Division, Washington, DC.

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3.10 PERIPHYTON DATA COLLECTION PROTOCOL

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Prepared by:

Jeffery Butt






2017

Disclaimer:

The mention of specific trade names or commercial products does not constitute

endorsement or recommendation for use.

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PURPOSE

The purpose of this document is to provide a standard protocol for DEP’s collection of

periphyton in lotic systems of Pennsylvania. The following sample and subsampling

types and protocols, laboratory procedures, as well as quality issues, will be discussed:

• Collection of Qualitative Multi-Habitat Periphyton samples

• Collection of Quantitative Benthic Epilithic Periphyton samples

• Subsample for Identification and Enumeration of Algal Taxa

• Subsample for the quantitative determination of Chlorophyll-a and Phycocyanin

Photopigments

• Subsample for the quantitative determination of Ash Free Dry Mass (AFDM)

• Subsample for the characterization of Cyanotoxins as part of qualitative multi-

habitat periphyton and benthic epilithic periphyton quantitative samples

• General laboratory considerations and procedures required to process the above

sample types

• Quality and harmonization issues concerning the identification and counting of

algae taxa.

INTRODUCTION

Characterizing lotic benthic algae communities is an important tool used by DEP in

determining the quality of its natural flowing waters. This is because, as is pointed out

by authors R. Jan Stevenson and Loren L Bahls,

“Benthic algae (periphyton or phytobenthos) are primary producers and an

important foundation of many stream food webs. These organisms also stabilize

substrata and serve as habitat for many other organisms. Because benthic algal

assemblages are attached to substrate, their characteristics are affected by

physical, chemical, and biological disturbances that occur in the stream reach

during the time which the assemblage developed (Barbour et al. 1999).”

In short, benthic algae and its analysis provides important information concerning the

condition of streams and rivers.

Algae, in this document, is a term used to describe a group of aquatic organisms that do

not represent a single formal taxonomic group. Rather, algae are defined as, in the

words of authors John D. Wehr and Robert G. Sheath, “… a loose group of organisms

that… [are] aquatic, photosynthetic, [and with] simple vegetative structures without a

vascular system…” (Wehr and Sheath 2003). Lotic periphyton includes several major

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taxonomic groups which, in Pennsylvania, are dominated by cyanobacteria, diatoms,

and green algae.

Benthic algae, in lotic systems, is collected by DEP as part of its on-going monitoring

responsibilities needed evaluate protection of surface water quality. Certain benthic

algae organisms, such as diatoms, are very useful because they have a very strong

relationship with nutrient concentrations and have been used as trophic indicators

(Potapova and Charles 2007, Porter et al. 2008, Hausmann et al. 2016).

Consequentially, collecting algae samples provides DEP with a tool that facilitates the

determination of trophic status and evaluate protection of surface water quality of rivers

and streams.

DEP biologists currently use two data collection procedures to collect periphyton from

lotic systems. These procedures are:

• Qualitative Multi-Habitat (QMH) Periphyton Data Collection Procedures

• Quantitative Benthic Epilithic (QBE) Periphyton Data Collection Procedures

The field procedures employed to collect both the QMH Periphyton and the QBE

Periphyton samples are appreciably derived from those described in the USGS

document Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as

Part of the National Water-Quality Assessment Program (USGS OFR 02-150, Moulton

et al. 2002). Many of the terms used in this DEP protocol are also found in this USGS

protocol and it is there that the reader may refer for definitions of these terms.

This document will also stipulate procedures and requirements that participating

laboratories must follow to consistently process and report results for the various

subsample types. Quality control practices needed to validate the identification and

enumeration of algal, cyanobacteria, and diatom taxa will also be discussed.

DEP biologists also collect water chemistry samples and discrete water parameters with

a properly calibrated field meter during algae sampling. Although a variety of chemical

analytes are often collected, which frequently depend upon the monitoring needs to

serve other objectives, the water chemistry that must be collected to support a

comprehensive analysis of the algae community should include Total Nitrate & Nitrite

Nitrogen, Total Nitrogen as N, Total Organic Carbon, Total Ortho Phosphorus as P,

Total Phosphorus as P, Dissolved Nitrate & Nitrite Nitrogen, Dissolved Ortho

Phosphorus, and Dissolved Phosphorus as P. Discrete water parameters collected with

a field meter should include Temperature, Specific Conductivity, pH, and Dissolved

Oxygen (both mg/l and % saturation).

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This document doesn’t address the data protocols for the collection and processing of

water chemistry samples or those for the collection of discrete water chemistry

parameters with a field meter. For these protocols, the reader is referred to Chapter 4 of

the Monitoring Book (Lookenbill and Whiteash 2021)

HISTORY OF LOTIC PERIPHYTON SAMPLING BY DEP

DEP began actively collecting lotic periphyton samples in 2005 at locations associated

with its Water Quality Network (WQN) sites. In 2012, biologists began collecting

samples for use in the development of a nutrient trophic index and concluded this

sampling in 2014. The samples collected for the nutrient trophic index have also been

applied to the development of an algae based Multi-Metric Index (MMI) for assessing

wadable and semi-wadeable Pennsylvania rivers and streams. DEP periphyton

sampling continues to this day for the purposes of developing this MMI as well as for

routine monitoring to document existing or changing biological conditions in rivers and

streams.

SAMPLING CONSIDERATIONS

Many different aquatic algae sampling options are available for the biologist to consider.

For example, the USGS OFR 02-150 lists four separate data collection protocols. These

include three quantitative protocols and one qualitative protocol, each of which are

designed to target different aquatic algae habitats. These USGS data collection

protocols include:

• Richest Targeted Habitat (RTH) – quantitative, represents a riffle or hard

substrate “… where the taxonomically richest algal or invertebrate community is

theoretically located.”

• Depositional Targeted Habitat (DTH) – quantitative, represents a depositional

habitat where fine sediments such as silt and sand accumulate.

• Phytoplankton (PHY) – quantitative, represents the water column habitat.

• Qualitative Multi-Habitat (QMH) – qualitative, whereas each of the three

quantitative sampling protocols defined above, in general, targets a single habitat

type, this sampling protocol is intended to represent the several different habitats

that may be present in a reach.

Currently DEP collects algae samples from lotic waters by using two different algae.

These two procedures include:

• QMH Periphyton Sampling Procedure (Similar to the USGS QMH procedure)

• QBE Periphyton Sampling Procedure (Similar to the USGS RTH procedure).

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The timing of algae sample collection is another important consideration for the

biologist. Algae growing in streams and rivers are readily influenced by high flow events,

temperature, and sunlight. Consequentially, the question of how long the biologist

should wait before sampling a stream or river after a high flow event must be

considered. Various researchers have studied the recovery of benthic algae

communities following a scouring flow and have made suggestions. For example,

Peterson and Stevenson (1990) suggested a three-week postponement in sampling

after a scour event to allow for recolonization (Peterson and Stevenson 1990, Barbour

1999). DEP biologists have considered the three-week “wait” period and, based on the

experience gained from observing the recovery of many streams in Pennsylvania, as

well as considering other logistical issues, have decided to use, in general, a two-week

“wait” period to resume sampling after a scour event. However, each stream or river

possesses its own scour response to flow which is based upon its own geomorphic

characteristic. The biologist must use knowledge of the watershed and best professional

judgement, as well as stream gauge data available on the USGS website (available at

https://waterwatch.usgs.gov/new/index.php?r=pa&id=ww current) to make decisions

regarding sampling delay after a scour event.

Temperature and sunlight availability also play a major role in the growth of algae in

rivers and streams. Typically, DEP biologists will begin sampling benthic algae in the

early spring in those streams that will later be shaded by the surrounding forest when

leaf-out occurs. Obviously, stream water temperatures are relatively cool in this early

spring-time period and will influence the composition of the algae community. Later,

biologists may return to the same stream to sample the algae community in mid-

summer and/or early fall when the stream is fully shaded and water temperatures are

more elevated so that they may characterize an algae community that may differ

appreciably from that observed in the spring. Sampling these streams across seasons

allows the biologist to better understand the transition in algae community composition

that occur with temperature and shading changes.

QMH PERIPHYTON SAMPLING PROCEDURE

The biologist may need to collect a QMH Periphyton Sample of microalgae and/or

macroalgae so that they may better understand the algal community present in any of

the multiple habitats available within a stream reach or area but not require area-based

(for area-based sampling, refer to the QBE Procedure discussed below) community

data, photopigment, or AFDM measures. This type of sampling aligns, in part, with the

USGS OFR 02-150 QMH sampling procedure. Because this procedure may sample

many types of algae habitats, there may be a cross over into the sampling situations

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referred to as DTH (Depositional Targeted Habitats) and RTH (Richest Targeted

Habitats) in the USGS OFR 02-150.

This sampling procedure provides DEP biologists with a tool to investigate special

monitoring questions. Such questions, for example, may address monitoring concerns

related to the habitat of early life phases of fish species or perhaps the algae community

composition that may lead to the production of cyanotoxin in non-thalweg stream

environments. In these examples, the sample location might be better described as a

specific habitat or area, such as a smallmouth bass young-of-the-year nursery habitat or

that of a small area where cyanobacteria are observed to be growing. Deciding to

delineate a sample reach versus a sample area is in part dictated by the question or

concern that the DEP biologist is attempting to address. Ultimately, the biologist must

characterize the habitat they sample and does so as described below in the data

collection section of this protocol.

With this protocol, the biologist should submit a subsample to a laboratory for algae

identification and enumeration. Also, the biologist should submit another subsample to a

laboratory to test for the presence of, cyanotoxins.

This protocol is intended for use in Pennsylvania wadeable riffle-run streams and semi-

wadeable large rivers. Refer also to Chapter 3.1, Wadeable Riffle-Run Stream

Macroinvertebrate Data Collection Protocol (Shull 2017b) and Chapter 3.4, Semi-

Wadeable Large River Macroinvertebrate Data Collection Protocol (Shull 2017a) for

terms and definitions that also apply to this section, as well as provide important safety

information for field staff to consider.

Field Data Sheet

The biologist will also complete the DEP QMH Periphyton Field Data Sheet (Appendix

A.7) at each monitoring site. To complete this field data sheet, the biologist must

provide information pertaining to the following (refer to the QMH Periphyton Field Data

Sheet in Appendix A.7 for more specific requirements for each data record):

• Site/sample information including water chemistry collection information and

personnel involved

• Latitude and longitude of the site

• Watershed Area – determined prior to making the field visit

• Field Meter Discrete Values – refer to Chapter 4.1, In-Situ Field Meter and

Transect Data Collection Protocol (Hoger 2020) for field meter use and

calibration requirements

• Total algae slurry volume (ml, minimum volume rules apply – refer to Table 1 in

the Subsampling Procedure section)

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• Densiometer and Compass Readings (refer to the Densiometer and Compass

Data Collection section – located near the end of the periphyton protocol - for

instructions in using these devices)

• Percent relative qualification of the periphyton habitat

• Notes pertaining to the three Algae ID Bottles collected

• Other notes (including any pertaining to cyanotoxin testing, if collected).

Habitat Types

To collect a QMH periphyton sample the biologist will need to employ several

approaches to sample across multiple habitats. The multiple habitats that may be

sampled will include any one or more of the following:

• Epilithic (Rock)

• Epiphytic (Plant)

• Epipelic (Silt)

• Epidendritic (Wood)

• Episammic (Sand)

• Gravel (subset of Epilithic material but listed independently here due to difference

in sampling protocol).

Data Collection

When sampling across multiple habitats, the biologist should proportionally represent

each habitat type in the targeted reach or area when making a composite sample. For

example, if after inspecting the targeted stream area, the biologist observes that the

multiple habitats present include about 45% Epiphytic, 10% Epipelic, 30% Epidendritic,

and 15% Episammic and no rock cobble or gravel (these percentages are recorded on

the field data sheet in the Periphyton Habitats section), then the final composited algae

slurry should be accumulated from a quantity of materials proportionally collected to

represent this habitat observation as best as possible. Because this protocol is

qualitative in nature the biologist must select sample material in a fashion that, in their

professional opinion, best reflects this observed habitat characterization. In other words,

based upon the above habitat characterization, the biologist would attempt to sample

material surfaces that roughly represents 45% Epiphytic, 10% Epipelic, 30%

Epidendritic, and 15% Episammic.

The multiple habitat sampling procedures, by type, will be performed as follows:

• For each sampling procedure described below, use de-ionized water to rinse the

scrubbed material into the algae slurry pan. The collector must remain aware of

the total amount of rinsate generated for the complete composited sample. In

short, the final accumulated algal slurry should be kept to a minimum by

conserving rinse water so that the total algae slurry (from all substrate) isn’t

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greater than that needed for lab analysis by a factor of two or three. This is

intended to maintain total slurry concentrations that better enable algae

identification and enumeration and cyanotoxin tests as well as facilitating

handling during sub-sampling homogenization.

• Epilithic (Rock) sampling performed by DEP is restricted to rock that is in the size

range of cobble (generally 2.5 to 10 inches) and is thus more easily moved by

the biologist. Boulders or bedrock are not readily transportable and are not

considered here. Gravel, which is a subset of epilithic, is treated separately in

this protocol because it is sampled differently. Cobble sampling is best

accomplished by removing cobble substrate from the benthos and transporting it

to the bank where the sampled surfaces may be scrubbed by a plastic or steel

brush while collecting the rinsate into the algae slurry pan. The sampled surface

should include scrubbing all that surface which is exposed to the aquatic

environment above the benthos.

• Epiphytic (Plant) sampling is best accomplished by clipping the aquatic plant,

using pruning shears or scissors, just above the sediment level (allowing roots to

remain in the sediment) and at the water line (retaining only the submerged

portion) and transporting the material to the bank where the total plant surface

may be scrubbed by a plastic brush. Collect the rinsate from this scrubbing into

the algae slurry pan. When scrubbing the plant surface, care should be taken so

that plant material is not inadvertently pulled off the plant and collected in the

pan. For plants that are less physically robust, such as grasses, the plants may

be placed in a capped plastic container, along with some de-ionized water, and

agitated vigorously. After agitating, the de-ionized water may be poured into the

collection pan and the plant material discarded. On the field data sheet, record

notes regarding the plant taxa from which samples were collected.

• Epipelic (Silt) sampling may be accomplished by using a silt/sand/gravel PVC

sampler and a kitchen spatula (see Figure 1). The silt/sand/gravel PVC sampler

may be constructed from a short section (about 3 or 4-inch length) of PVC pipe

(4 or 5-inch diameter) and capped at one end. After placing this PVC sampler in

the water, rotate it up-side-down to remove air. Once the air is removed, push the

open end of the PVC sampler a few millimeters into the silt. Without moving the

PVC sampler, slide the plastic kitchen spatula underneath the PVC sampler to

secure the upper layer of silt inside the PVC sampler. Carefully extract the PVC

sampler and spatula from the stream without spilling the contents. Pour the

contents of the PVC sampler into the collection pan. As with all collecting

strategies used for this QMH procedure, the final accumulated algal slurry should

be kept to a minimum so that the total algae slurry (from all substrate) isn’t

greater than that needed for lab analysis by a factor of two or three. Quantities of

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homogenized algal slurry needed to perform each laboratory test are indicated in

Table 1.

Figure 1. Silt/sand/gravel PVC and spatula sampling apparatus. This apparatus

may consist of a section of capped 4 or 5-inch diameter PVC pipe and a kitchen

spatula. The spatula must be large enough so that when it is slid under the PVC

pipe it will prevent the loss of silt, sand, or gravel from the sample as it is pulled

from the benthos.

• Epidendritic (Wood) sampling may be best accomplished by collecting

submerged wood and transporting it to the bank where the surfaces may be

scrubbed with a plastic or metal brush while collecting the rinsate into the algae

slurry pan. Use de-ionized water to wash the scrubbed material into the algae

slurry pan. When scrubbing the wood surface, care should be taken so that wood

material is not inadvertently pulled off and collected in the pan. The biologist may

find it useful to use a small saw to cut portions of larger submerged branches for

removal to the scrubbing area. As with all collecting strategies used for this QMH

procedure, the final accumulated algal slurry should be kept to a minimum by

conserving rinse water so that the total algae slurry (from all substrate) isn’t

greater than that needed for lab analysis by a factor of two or three. Quantities of

homogenized algal slurry needed to perform each laboratory test are indicated in

Table 1.

• Episammic (Sand) sampling is accomplished in much the same way as that

described for Epipelic (Silt) by using a silt/sand/gravel PVC sampler and a plastic

kitchen spatula (see Figure 1). The silt/sand/gravel PVC sampler may be

constructed from a short section (about 3 or 4-inch length) of PVC pipe (4 or 5-

inch diameter) and capped at one end. After placing this PVC sampler in the

water, rotate it up-side-down to remove air. Once the air is removed push the

open end of the PVC sampler a few millimeters into the sand. Without moving the

PVC sampler, slide the plastic kitchen spatula underneath the PVC sampler to

secure the upper layer of sand inside the PVC sampler. Carefully extract the

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PVC sampler and spatula from the stream without spilling the contents. Pour the

contents of the PVC sampler into a capped plastic bottle and agitate vigorously.

After allowing the sand to settle to the bottom of the container pour the agitate

(with algae entrained) into the collection pan. As with all collecting strategies

used for this QMH procedure, the final accumulated algal slurry should be kept to

a minimum so that the total algae slurry (from all substrate) isn’t greater than that

needed for lab analysis by a factor of two or three. Quantities of homogenized

algal slurry needed to perform each laboratory test are indicated in Table 1.

• Gravel sampling is accomplished in the same way as that described for

Episammic (Sand). Refer to the Episammic (Sand) sampling procedure provided

above.

Subsampling Procedure

QMH Periphyton samples must always be subsampled for algae identification and

enumeration and sometimes, depending on the requirements of the project, cyanotoxin

analysis. Subsampling may be performed in the field if electric is available to operate a

blender and a magnetic stirrer. If subsampling can’t be performed in the field the sample

may be temporarily stored in labeled 500ml bottles that are capped and stored in a wet

ice cooler to await later treatment.

Note: This is an important step needed to prevent cross-contamination between

samples. Prior to performing any subsampling procedures on a site sample, thoroughly

clean all equipment that may come in contact with the algae slurry sample and, at the

conclusion of this cleaning, rinse with de-ionized water. Because site samples are often

accumulated through the field day and subsampled at the conclusion of that day’s work,

all equipment must be appropriated cleaned between samples.

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Table 1. QMH Periphyton Subsample Tests: Homogenized Volume (ml), Bottleware,

Preservation, Transportation, and Storage

Subsample Test Homogenized

Volume (ml) Bottleware Preservation Transportation Storage

Algae ID Bottle #1 100 120ml

Plastic

7ml

Formaldehyde Cool, dry, dark

Cool, dry,

dark


Plastic

7ml


Cool, dry,

dark


Plastic

7ml


Cool, dry,

dark

Minimum Algae Slurry Volume

(plus excess) without

Cyanotoxin Test

400

Cyanotoxin Test 250 Whirl-Pak® None Dry-ice -80°C


(plus excess) with Cyanotoxin

Test

700

Subsampling consists of the following steps:

Slurry Preparation

Once all algae slurry is collected, pour the contents of the collection pan into a large

plastic beaker (beaker should be large enough to contain at least 2000 ml of algae

slurry). Dilute the algae slurry with de-ionized water to provide enough material for all

subsample tests (refer to Table 1 for minimum quantities required). Since this procedure

is intended to be qualitative in nature, sample surface area is not determined and the

ID/enumeration laboratory will not report results in the same units as that provided for

quantitative samples (see below). Consequentially, the quantity of surface area sampled

or quantity of algae slurry collected is not reported to the laboratory.

Further, the identification/enumeration QA process requires the submission of blind

duplicates to laboratories. Typically, identification/enumeration blind duplicates are

submitted at a quantity approximately equal to 10% of the total quantity of samples

submitted to the laboratory. Consequentially, multiple ID/Enumeration bottles are

collected from each sample.

Subsampling for Algae Identification/Enumeration and Cyanotoxin

Once the dilution process (if required to achieve minimum quantities stipulated in Table

1) is complete and the final algae slurry sample volume is recorded, the sample must be

homogenized. Often a handheld low-speed blender is applied to the contents of the

beaker to break apart the larger pieces of algae. This blending also helps to better

homogenize the algae slurry. A high-speed blender shouldn’t be used at this point

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because at high speeds the blender may lyse the algae cells, making the identifications

more difficult.

Once blended, the algae slurry beaker may then be placed on a magnetic stirrer to be

maintained in a homogenized state as subsamples are extracted (see Figure 4). Pour

100ml of well homogenized algae slurry into a graduated cylinder to ensure accuracy of

volume measurement – then pour this volume into a labeled 120ml algae

ID/enumeration sample bottle being careful to leave no algae residue in the graduated

cylinder. Return the slurry beaker to the magnetic stirrer to resume homogenization

prior to pouring the second and third 100ml algae ID/enumeration subsamples.

Preserve each identification and enumerations subsample with 7ml of formaldehyde

(volume of formaldehyde preservative must be reported to the ID/enumeration

laboratory). Refer also to Table 1 for subsample and preservative volumes as well as for

transportation and storage requirements.

To subsample for Cyanotoxin return the algae slurry beaker to the magnetic stirrer to re-

homogenize. Pour 250ml of homogenized sample into a Whirl-Pak® bag and seal. Then

transport and store as prescribed in Table 1.

Labels and Submission

Algae Identification and Enumeration bottles will be labeled with a GIS Key (e.g.,

20170516-1115-jbutt, indicating the date, time, and collector name), DEP Algae

Sampling Site ID Number, Stream/River Name, and Location Description.

An example of an Algae Identification and Enumeration Laboratory Submission Data

sheet can be found in Appendix A.8. This data sheet provides essential information

required by the laboratory performing identifications and is used to determine final

reporting values to DEP. Also, location information is provided on this submission sheet.

This location information may assist the laboratory performing identifications in

providing more accurate algae taxa identifications, especially as they may pertain to

species or variation level identifications of diatoms.

QBE PERIPHYTON SAMPLING PROCEDURE

Benthic epilithic periphyton is the preferred sampling substrate targeted by DEP to

collect data pertaining to lotic algal communities for use in its monitoring program. One

reason for this preference is that many streams and rivers in Pennsylvania are relatively

fast-flowing and have a geomorphology that creates riffles where cobble substrate is

plentiful. These cobbled riffles present the DEP biologist with a consistent substrate

from which to extract samples across many streams in Pennsylvania. Consistency in

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sample substrate is needed to support the diatom MMI development and

implementation. As noted by Barbour et al., “For comparability of results, the same

substrate/habitat combination should be sampled in all reference and test streams”

(Barbour et al. 1999). Also, as described by Kelly et al., “Under many circumstances

(particularly at fast-flowing sites), the epilithon is the most abundant substratum,

sampling procedures are relatively straight-forward and ecological preferences of most

common species are well understood” (Kelly et al. 1998). Kelly et al. also points out

that, regarding the epilithon, “The performance of major diatom-based indices on this

substratum is well understood” (Kelly et al. 1998).

In 2005, Marina Potapova and Donald Charles analyzed the large algae data set

created by the USGS National Water-Quality Assessment Program (USGS NAWQA,

Potapova and Charles 2005) and demonstrated several important findings regarding the

type of substrate sampled. The USGS NAWQA program collected as many as four

different sample types and compared results from paired samples from hard and soft

substrates. From this analysis, those findings that support the use of the benthic epilithic

periphyton, given the typically good availability of cobble substrate in many

Pennsylvania streams, are:

• “Ordinations of assemblages from hard and soft substrates were highly

concordant and provided similar information on environmental gradients

underlying species patterns.”

• “…only one sample has to be collected at each site for water quality assessment

surveys, and that sample can come from whatever substrate is available.”

Consequentially, based upon the recommendations from the scientific literature, the

nature of the monitoring activity pursued by DEP, and the characteristic of the streams

and rivers typically being sampled in Pennsylvania, DEP chose and continues to use a

benthic epilithic periphyton sampling protocol. This protocol is intended for use in

Pennsylvania wadeable riffle-run streams and semi-wadeable large rivers.

Site Selection

Within the area where the stream is to be sampled, select a reach in which a riffle is

located and which is, in general, representative of the stream in that locale.

Representative feature considerations should include wetted width, substrate type and

size, shading, and standing periphyton crop.

Transects and Rock Selection

Rocks must be extracted randomly from the stream or river along three transects at

each sampling site. The term transects, as it is used here, should not be confused with

the same term as it is used in Chapter 4.1, In-Situ Field Meter and Transect Data

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Collection Protocol (Hoger 2020). Regarding periphyton sample collection, these

transects are only used to facilitate the random selection of rocks as demonstrated in

Figure 3 and to measure wetted width of the stream for data recording on the field data

sheet.

Rocks (sometimes referred to as cobble) to be extracted for sampling must be of the

size and shape to facilitate the rock scrub. A suitable sample rock is one that allows the

foam ring on the PEP Sampler to seal properly (refer to the section “Collecting the

Periphyton Slurry – The Rock Scrub,” located below, for a description of the PEP

Sampler and refer to Figure 4). For the foam ring to seal properly, the sample rock must

be small enough to fit in the PEP Sampler but be larger than the approximately 3-inch

diameter foam gasket. Also, the rock must be relatively smooth (flat) so that ridges or

other irregularities in the scrub surface area do not interfere with the foam gasket seal

and allow the use of the scrubbing and slurry collection tools. If the first rock picked at

each location will likely not facilitate the scrub process, choose the next suitable rock

located immediately upstream at that same transect location. As with all other aspects

of this protocol, experience will be the best guide for the biologist in making appropriate

choices in selecting appropriate sample rocks.

The quantity of rocks that must be sampled at each site varies based upon the size of

the watershed located upstream of the site location. Watersheds with areas equal to or

greater than 1,000 square miles (2,590 square kilometers) are to be considered as a

semi-wadeable large river and will require a 27-rock QBE periphyton sample to properly

represent the habitat and species variability in that sample reach. Watersheds smaller

than 1,000 square miles in area should be considered as a wadeable riffle-run stream

that will require a 9-rock sample. Watershed area is an attribute that must be

determined through a GIS based inquiry which must be performed prior to sampling in

the field. For example, Pine Creek in Lycoming County, immediately upstream of the

confluence with Ramsey Run at N41.283650 W77.320917, has an upstream watershed

area of 940 square miles and is classified as a wadeable riffle-run stream thus requiring

a 9-rock sample. Whereas, the West Branch Susquehanna River at Lewisburg,

downstream of the Route 45 bridge at N40.9653 W76.8782, has an upstream

watershed area of 6,820 square miles and is considered a semi-wadeable large river

requiring a 27-rock sample. Refer to Chapter 3.1, Wadeable Riffle-Run Stream

Macroinvertebrate Data Collection Protocol (Shull 2017b) and Chapter 3.4, Semi-

Wadeable Large River Macroinvertebrate Data Collection Protocol (Shull 2017a) for a

more complete definition of these types of surface water characterizations as well as

important safety procedures that must be considered by field staff before working in

these waterbodies.

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Setting up Transects and Making the Sample Rock Selections in Wadeable Riffle-Run

Streams

To facilitate the random rock selection in wadeable riffle-run streams, the biologist must

choose a sample reach that longitudinally (parallel to the stream thalweg) measures no

more than about 100 meters in length and in which is located enough riffle to deploy the

three sample transects – each transect must be deployed over a riffle. An example of

such a sample reach is shown in Figure 2.

Figure 2. An example of a periphyton sample reach in a wadeable riffle-run stream.

Three transect measuring tapes are pulled across the riffle section of the stream to

facilitate random rock selections.

The step-by-step process for setting the three transect lines, computing the random

rock sample pick locations, and collecting the rocks is as follows:

1. Three measuring tapes must be pulled across the stream to create three

transects. Each tape is pulled across the stream so that it is oriented

perpendicular to the direction of flow in the stream in a manner like that shown in

Figure 2.

2. The biologist must exercise care when pulling the transect tapes across the

stream so that they do not step on, or interfere with, the substrate that will be

targeted for sampling. In general, the collector should remain within a few feet of

the tape on the downstream side, thereby leaving substrate on the upstream side

of the tape undisturbed and available for sampling. Also, to lessen the chance of

disturbing the substrate along any transect, the biologist should not closely

situate any of the transect tapes to one-another. Widely spacing the transect

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tapes over the sample reach helps to prevent biologists from disturbing substrate

in adjacent transects and additionally helps the biologist to better represent the

potential habitat variation in that stream.

3. Transect tapes should be pulled beyond the wetted margins of the stream so that

wetted width may be accurately measured. The wetted stream width parameter is

used to calculate random rock selection locations.

4. Record the wetted margin tape measurements for each transect at the RDB

(Right Descending Bank) and the LDB (Left Descending Bank) locations on the

QBE Periphyton Field Data Sheet. In streams to wide for pulling a tape, the

biologist should use a laser range finder. An example of this record, along with a

transect random rock pick calculation, is provided in Figure 3. This figure is an

excerpt from the DEP QBE Periphyton Field Data Sheet (Appendix A.6).

5. Determine the wetted width for each transect and record this width in the QBE

Periphyton Field Data Sheet.

6. Divide the wetted width for each transect by 3. The result of this mathematical

operation is referred to as the 1/3rd transect distance. Record this 1/3rd distance

for each transect on the data sheet. Essentially, each transect is divided into

equal thirds and, for a 9-rock sample (wadeable riffle-run stream), a single rock is

extracted from a randomly determined location in each of the 1/3rd transect

sections. In a 27-rock semi-wadeable sample, three rocks will be extracted from

randomly determined locations in each of the 1/3rd transect sections.

7. Random percentage numbers are used to compute sample locations along the

tape in each of the 1/3rd transect sections. Random percentage numbers may be

taken from a random number table or from the roll of a 10-sided die (the first die

roll represents the tens place of a percentage use in the location calculation – for

example, a roll of 1 represents 10. The second die roll represents the units place

– for example, a roll of 5, when combined with the previous die roll, will be

recorded as the random percentage of 15%). Record three random percentages

– which may range from 0% to 99% - for each 1/3rd transect section in the

designated spaces of the field data sheet.

8. Compute the tape locations on each transect to determine where random picks

are to occur. For example, as shown in Figure 3 below, the rock pick location on

Transect 1 in the first 1/3rd location would be computed as: (15% x 16.3ft) + 3.8 =

6.2ft on the tape measure. In this example, 16.3ft is the 1/3rd distance on transect

1 tape and 3.8ft is the RDB location. The second rock pick location on Transect

1, located in the second 1/3rd section would be computed as: (78% x 16.3ft) +

16.3ft (first 1/3rd section) + 3.8ft (the RDB) = 32.8ft on the tape measure. The

remaining rock pick locations for this 9-rock sample are similarly computed.

Locations for a 27-rock semi-wadable sample are computed in a similar fashion

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except each 1/3rd transect section will use three random percentage numbers

instead of one random percentage number.

9. Once the random sample locations along each transect are computed, the

biologist must collect suitable sample rocks from each location. Rock samples

must be collected from upstream of the transect tape and the biologist should

collect the first suitable sample rock encountered when moving upstream from

the transect tape location. As previously discussed, a suitable sample rock is one

with the characteristics that allows the foam ring on the PEP Sampler to seal

properly. Refer to the section “Collecting the Periphyton Slurry – The Rock

Scrub,” located below, for a description of the PEP Sampler and refer to Figure 4.

If the first rock picked at each location will likely not facilitate the scrub process,

choose the next suitable rock located immediately upstream at that same

transect location.

10. When collecting the sample rocks, the biologist should prevent any contact with

the surface of the rock exposed to the stream water – in other words, lift the rock

by touching it only from the embedded surface (bottom) side. Collect the sample

rocks into a plastic bin, being careful to avoid any contact of the upper surface of

the rock with the container or other rocks. This care is needed because even the

slightest disturbance to the sample area on the rock will skew the results

generated during the laboratory subsample analyses. (refer to the section

“Collecting the Periphyton Slurry” for a description of that sample area)

11. Transport the sample rocks to the field sample processing station and prepare for

the collection of the Periphyton Slurry. Collection of the Periphyton Slurry is

described below. The field sample processing station, as shown in Figure 4,

should be setup in a fashion that allows for the positioning of all field sampling

hardware on a horizontal surface, such as on a table or the foredeck of a boat,

and located where biologists may process the sample in a manner that minimizes

the likelihood of fouling or loss of sample material due to environmental elements

such as rain or wind, for example.

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Figure 3. Excerpt from the DEP QBE Periphyton Datasheet demonstrating the random

rock pick calculation made for a hypothetical small stream (wadeable riffle-run) that

requires a 9-rock sample.

Setting up Transects and Making the Sample Rock Selections in Semi-Wadeable Large

Rivers

To facilitate the random rock selection in semi-wadeable large rivers, the biologist must

choose a sample reach that longitudinally (parallel to the river thalweg) measures no

more than about 100 meters in length and in which is located enough riffle to deploy the

three sample transects – each transect must be deployed over a riffle.

Sampling in semi-wadable large rivers often differs from that of sampling in wadeable

riffle-run streams in four important ways. One difference may be the wetted width of the

river – it may be too wide for the positioning of a measuring tape. In these wide wetted-

width locations a laser range finder should be used instead of a tape measure. A

second important difference relates to depth – in such rivers, the biologist may

encounter sections of the transect that are too deep to sample. A section of river is too

deep to sample if the biologist is unable to safely grab substrate from the benthos while

wearing waders. If an area of the transect is too deep for sampling, the biologist must

bypass that area and collect additional sample rocks from those sections of the transect

that may be safely sampled and note this adjustment on the QBE Periphyton Field Data

Sheet. A third difference is that the biologist should expect to facilitate the sampling by

using a power boat, canoe, or kayak. The last difference relates to the possibility of

multiple sampling zones, as defined by water influence transitions, occurring at a river

site. Situations involving multiple sampling zones, and how to handle them, are

discussed in the next paragraph.

Multiple sampling zones may need to be created to properly sample periphyton in semi-

wadeable large rivers. This occurs when field meter transect data indicates a relative

lack of homogeneity in parameters such as temperature, pH, specific conductivity,

Transect 1

RDB LDB W / 1/3

3.8 ft 52.7 ft 48.9 / 16.3

Transect 2

RDB LDB W / 1/3

7.8 ft 49.6 ft 41.8 / 13.9

Transect 3

RDB LDB W / 1/3

14.2 ft 63.1 ft 48.9 / 16.3

Random #'s: 89 Random #'s: 72

Transect Random Rock Pick Calculation

(Upstream) Random #'s: 15 Random #'s: 78 Random #'s: 25

6.2 ft 32.8 ft 40.5 ft

(Middle) Random #'s: 33 Random #'s: 42 Random #'s: 11

15.0 ft 45.0 ft 58.5 ft

12.4 ft 27.5 ft 37.1 ft

(Downstream) Random #'s: 5

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dissolved oxygen, or turbidity when measured along a river transect and is

characteristic of water influences originating from un-mixed tributary influences. When

such water quality heterogeneity is observed, a semi-wadeable river will need to be

subdivided into separate sampling zones and a 27-rock periphyton sample be

independently collected from each zone of influence. For example, the Susquehanna

River near Harrisburg, based on transect data, shows three distinct zones of water

influence, these being the “East” zone, the “Middle” zone, and the “West” zone. When a

river must be subdivided into multiple zones, transect sets in each zone must be

restricted to the width of river section that defines that zone.

Once these differences are accounted for, the step-by-step process for setting the three

transect lines, computing the random rock pick locations, and collecting the rocks is

identical to that defined above for a 9-rock wadeable riffle-run stream except that a 27-

rock sample is used.

Regarding the use of a boats, the biologist should familiarize themselves with the

targeted site prior to going to the field to sample. The target sample locations may be

located some distance from points of access and will thus require the use of a power

boat to facilitate the field work. In some instances, all that is needed is a canoe or a

kayak. The biologist is reminded that all Pennsylvania Fish and Boat Commission

boating laws and safe boater guidance must be observed if sampling with a boat.

Collecting the Periphyton Slurry – The Rock Scrub

After the rocks are transported to the field sample processing station location, the

biologist will scrub the rocks to collect the periphyton slurry. This slurry collection is

made using a Pennsylvania Epilithic Periphyton (PEP) Sampler, de-ionized water, scrub

brushes, pipets, squirt bottles, and collection containers (500ml plastic bottles, refer to

Figure 4). The PEP Sampler is a clamping device that secures a rock sample and

provides a fixed circular sample area (equal to 20.6cm2) and a foam gasket seal (toilet

seal) that is clamped to rock (cobble) substrates using partially flexible clamping boards

that can be tightened with quick release clamps. The following procedure steps will be

used to conduct the rock scrubs:

1. After clamping the rock in the PEP Sampler, with the top face of the rock facing

up, verify that the foam ring seal is secure by squirting an adequate amount of

de-ionized water into the scrub chamber and observing that no water leaks out. If

no leaks are observed, continue to step 2, otherwise re-clamp the rock in the

PEP Sampler and re-verify that the seal does not leak. As previously discussed,

for the foam ring to seal properly the rock must be small enough to fit in the PEP

Sampler but be larger than the approximately 3-inch diameter foam gasket. Also,

the rock must be relatively smooth (flat) so that ridges or other irregularities in the

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scrub surface area do not interfere with the foam gasket seal and allow the use

of the scrubbing and slurry collection tools.

2. Using the scrub brushes and a small amount of de-ionized water, scrub the rock

area within the PEP sampler. After a liberal amount of scrubbing, pipet the algae

slurry out of the PEP sampler and into the 500ml sample collection bottle(s).

Repeat this process until the rock surface area within the PEP sampler is clean –

the rock surface is clean when the de-ionized water is relatively clear after

scrubbing. Determining when this process is complete and the scrub water is

clear, is a matter of professional judgement and experience because, depending

on the composition of the rock, continued scrubbing may displace mineral

rendering the scrub water unclear. When scrubbing is complete, unclamp the

rock and dispose of the rock. However, if stream site is to be repeatedly sampled

and cobble is scarce, the biologist may choose to return the cobble to the riffle so

that sampleable substrate is not depleted.

At this point, it is important to note that because the periphyton community is a

three-dimensional arrangement of often microscopic organisms, sometimes

stratified into layers, all of which contribute to the base of the food web within the

stream and may strongly influence the cascade of energy and nutrients through

all trophic levels in the stream, it is important for the biologist to accurately

represent that community through the diligent collection of algae material.

3. Repeat steps 1 and 2 until all rocks have been scrubbed and all algae slurry is

collected into the 500ml collection bottles. Sometimes multiple 500ml collection

bottles will need to be utilized for a single sample. All 500ml bottles used to

collect the algae slurry at a site should be labeled and the quantity of bottles

noted in a comments field for the QBE Periphyton Field Data Sheet – this is to

ensure that the biologist, when processing subsamples at the end of the

sampling day, doesn’t forget to include all sample bottles when compositing the

algae slurry.

4. Cap the 500ml collection bottles, being careful not to spill any of the slurry. Label

the bottles with the date and site information and place in a dark cooler with wet

ice to await the subsampling procedure – do not freeze.

5. Clean all equipment including the PEP sampler, brushes, etc. First scrub all

equipment in the stream water followed by a rinse with de-ionized water. Through

cleaning is required to prevent the contamination of samples with residue from

other sites.

6. Organize and stow all gear for safe transport to the next work site.

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Figure 4. The Pennsylvania Epilithic Periphyton (PEP) Sampler with various tools used

to sample algae from rock substrate. Rocks suited for periphyton scrubbing must be of

size and shape to fit properly in the PEP sampler – large enough and flat enough for the

sampler circular foam gasket to form a leak-proof seal around the area to be scrubbed.

Field Data Sheet

The biologist will also complete the DEP QBE Periphyton Field Data Sheet at each

sampling location (Appendix A.6). In addition to performing the Transect Random Rock

Pick Calculation as discussed above, the biologist must provide information pertaining

to the following (refer to the QBE Periphyton Field Data Sheet for more specific

requirements for each data record):

• Site/Sample Information including water chemistry collection information and

personnel involved

• Latitude and longitude of the site

• Watershed Area – determined prior to making the field visit

• Representation of the Inorganic Substrate

• In-situ Field Meter Discrete Values – refer to the Field Meter and Transect Data

Collection Protocol Section in Chapter 4 for field meter use and calibration

requirements

• Densiometer and Compass Readings (refer to the Densiometer and Compass

section of this document for instructions for use these devices)

• Quantity of rocks sampled (equates to surface area scrubbed – each rock =

20.6cm2)

• Transect random rock pick calculations

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• Total algae slurry volume (ml, minimum volume rules apply – refer to

Subsampling Procedure section)

• Notes pertaining to the Algae ID Bottles collected

• Notes, including collector and sequence numbers, pertaining to the Phycocyanin

sample bottle

• Notes, including collector and sequence numbers, pertaining to the Acidified

Chlorophyll-a filters

• Notes, including collector and sequence numbers, pertaining to the AFDM filters

• Other notes (including any pertaining to cyanotoxin testing, if collected).

Subsampling Procedure

The QBE Periphyton Sampling procedure requires subsampling for the identification

and enumeration of algal taxa, the quantitative determination of chlorophyll-a and

phycocyanin photopigments, the quantitative determination of AFDM, and in some

cases for the characterization of cyanotoxins. This subsampling may be performed in

the field at the sample site (if electricity is available for operating a blender and a

magnetic stirrer) or the sample may be temporarily stored in labeled 500ml bottles that

are capped and stored in a wet ice cooler to await later (same day) subsampling

treatment. Table 2 provides information regarding test volumes, bottle or filter

requirements, preservation procedure, as well as transportation and storage needs for

the QBE subsamples.

Note: This is an important step needed to prevent cross-contamination between

samples. Prior to performing any subsampling procedures on a site sample, thoroughly

clean all equipment that may come in contact with the algae slurry sample and, at the

conclusion of this cleaning, rinse with de-ionized water. Because site samples are often

accumulated through the field day and subsampled at the conclusion of that day’s work,

all equipment must be appropriated cleaned between samples.

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Table 2. QBE Periphyton Subsample Tests: Homogenized Volume (ml), Bottleware,

Preservation, Transportation, and Storage

Subsample Tests Homogenized

Volume (ml)

Bottle or

Filter Preservation Transportation Storage


Plastic

7ml


Cool, dry,

dark


Plastic

7ml


Cool, dry,

dark


Plastic

7ml


Cool, dry,

dark

Chl-a Filter #1 (Acidified) 2 WhatmanTM

GF/F

1ml MgCO3 -

Fold-Foil wrap Dry-ice -80°C


GF/F

1ml MgCO3 -



GF/F

1ml MgCO3 -


Phycocyanin Bottle 30 120ml

plastic None Wet-ice Refrigerate

AFDM Filter #1 2 WhatmanTM

GF/F Fold-Foil wrap Dry-ice -80°C






(plus excess) without

Cyanotoxin Test

400

Cyanotoxin Test 250 Whirl-Pak® None Dry-ice -80°C


(plus excess) with Cyanotoxin

Test

700

Subsampling consists of the following procedures and must be done in the order listed:

Initial Slurry Preparation

Composite the contents of all the 500ml collection bottle(s) for a site into a large plastic

beaker (beaker should be large enough to contain at least 2000 ml of algae slurry, or

more, and is able to be used with the magnetic stirrer and low-speed blender). Once all

slurry is collected into the beaker, apply a low-speed hand-held blender to this slurry.

This blending helps to homogenize the slurry by breaking apart the larger pieces of

algae. It is important that a high-speed blender with razor-sharp blades not be applied

at this point. The algae identification/enumeration bottles must be poured off prior to the

application of high-speed blending with razor-sharp blades – such blending will lyse

algae cells, making identifications and counting more difficult.

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Once this initial blending is complete in the 2000ml beaker, pour the complete algae

sample into multiple 1000ml graduated cylinders – as many as is required to contain the

entire sample. Next, dilute the algae slurry with de-ionized water to provide enough

material for all subsample tests (refer to Table 2 for minimum quantities required). For

example, if the biologist is electing not to subsample for the cyanotoxin test, but is

intending to subsample for ID/Enumeration, Chlorophyll-a, Phycocyanin, and AFDM

only, then the minimum quantity of algae slurry needed is 400ml. However, if the

biologist is electing to subsample for cyanotoxin, in addition to all the other subsamples,

then the minimum quantity of algae slurry needed is 700ml. These minimum quantities

will insure that enough algae slurry is available for all subsamples plus a little extra (in

case of small spills and enough material being available for the magnetic stirrer to work

and the Hensen-Stemple extractions to be made). The biologist should not over dilute

the algae slurry – the laboratory tests are best facilitated by providing as concentrated a

subsample as possible. If after compositing all the 500ml collection bottles for a site, the

total algae slurry is greater than the minimum quantity required, do not further dilute

beyond that needed to move the volume to the next highest 100ml increment level – for

example, if after compositing all bottles, the total volume is found to be about 1,244ml,

then dilute to 1,300ml. Doing this helps to insure accuracy of volume measurement in a

1000ml graduated cylinder which are often incremented in units of 100ml.

The total algae slurry volume must be recorded on the field data sheet and, eventually,

reported to the laboratory performing the identifications/enumerations. Since the

ID/enumeration laboratory must provide results to DEP that enable the biologist to

convert results to cells/cm2 and report biovolume (unit of µm3/cm2) estimates, accurate

reporting of algae slurry volume and surface area scrubbed is essential. Also note that

the identification/enumeration QA process requires the submission of blind duplicates to

laboratories. Typically, identification/enumeration blind duplicates are submitted at a

quantity approximately equal to 10% of the total quantity of samples submitted to the

laboratory. This is why multiple ID/Enumeration bottles are collected from each sample.

This QA process is discussed in detail later in this protocol.

Subsampling for Algae Identification/Enumeration

Once the Initial Slurry Preparation process is complete, the first subsampling that must

be completed is that needed for the Algae Identification and Enumeration.

Pour all graduated cylinders from the Initial Slurry Preparation Process back into the

2000ml beaker. This total sample is then homogenized on a magnetic stirrer (see Figure

5). Once the sample is well homogenized (this may take approximately 30 seconds on

the magnetic stirrer and is evident when a good vortex is observed to be established in

the beaker), pour 100ml of the homogenized algae slurry into a 100ml graduated

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cylinder to insure accuracy of volume measurement. Thereafter, pour this volume into a

labeled 120ml algae ID/enumeration sample bottle while being careful to leave no algae

residue in the graduated cylinder. Do not use de-ionized water to rinse this residue into

the ID/enumeration bottles – adding de-ionized water at this point will further dilute the

sample and introduce volume related discrepancies into the final results reporting. A

swirling motion on the 100ml graduated cylinder followed by a quick pour into the

sample bottle will usually prevent sample material from being retained in the graduated

cylinder and will insure that all material is transferred. Sometimes a small quantity of

sample material must be returned to the graduated cylinder from the sample bottle and

a re-pour used to transfer all algae material. After collecting an ID/enumeration bottle,

return the slurry beaker to the magnetic stirrer to re-establish homogenization prior to

pouring the second 100ml algae ID/enumeration sample and later, the third 100ml algae

ID/enumeration subsample.

Preserve each 100ml ID/enumeration subsample with 7ml of formaldehyde (volume of

formaldehyde preservative must also be reported to the ID/enumeration laboratory).

Refer also to Table 2 for preservative volumes as well as for transportation and storage

requirements.

Algae Identification and Enumeration bottles will be labeled with a GIS Key (e.g.,

20170516-1115-jbutt, indicating the date, time, and collector name), Stream/River

Name, and Location Description.

Prior to submitting the algae ID samples to a laboratory, an Algae Identification and

Enumeration Laboratory Submission Data sheet must be made and submitted to the

laboratory along with the samples. An example of such a submission sheet can be

found in Appendix A.8. This data sheet provides essential information required by the

laboratory performing identifications and is used to determine final reporting values to

DEP. Also, sample site location information is provided on this submission sheet. This

location information may assist the laboratory performing identifications in providing

more accurate algae taxa identifications, especially as they may pertain to species or

variation level identifications of diatoms.

Note that algae ID sample bottles, once preserved, may be stored in a cool, dry, and

dark location (refer to Table 2), for upwards of several months prior to sending to an

identification lab. Often, DEP will retain these samples until the end of a collection

season prior to shipping all samples together to the identification/enumeration

laboratory.

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When ID/Enumeration subsampling is complete, the biologist may then proceed to

subsampling for Phycocyanin, Acidified Chlorophyll-a, AFDM, and Cyanotoxins.

Subsampling for Phycocyanin, Acidified Chlorophyll-a, and AFDM

After the Algae Identification/Enumeration subsampling has been completed, the slurry

may be further blended in a highspeed blender. Highspeed blending in a food-grade

processing type of blender (those with stainless steel “razor sharp” blender blades) is

needed to properly prepare the algae for phycocyanin, acidified chlorophyll-a, and

AFDM subsampling in samples with robust filamentous or other forms of algae that

resisted the low-speed blending. When highspeed blending is complete, return the

sample to the 2000ml beaker and magnetic stirrer to provide continuous

homogenization.

Hereafter, the order of subsampling will first be phycocyanin, followed by AFDM, then

acidified chlorophyll-a, and lastly, if needed, cyanotoxin.

At present, all phycocyanin, acidified chlorophyll-a and AFDM subsamples are delivered

to DEP Bureau of Laboratories (BOL) for further processing. These samples may either

be hand delivered to BOL or may be shipped to BOL by courier. Refer to Table 2 for

methods of transportation and storage. All necessary laboratory submission sheets

must be completed and delivered to BOL along with the sample delivery. Phycocyanin

has a very short holding time for pre-processing at BOL and should either be hand-

delivered to BOL at the end of the field day or delivered overnight by courier.

Replicate samples for Phycocyanin, Acidified Chlorophyll-a, and AFDM

Although appreciable effort is made by DEP biologists to thoroughly homogenize the

algae slurry, variation in duplicate subsample results have been noted in nearly all

reported lab results. This variation is, in part, due to robust forms of filamentous algae

that resist complete homogenization, even after the use of a highspeed food-grade

blender. The best way to deal with this unavoidable variation in photopigment and

AFDM results is to run replicate subsamples. What has proven to be effective is

collecting three replicate subsamples each for phycocyanin, acidified chlorophyll-a, and

AFDM tests. The biologist may then utilize the average, median, minimum, and

maximum values generated by the three replicate subsamples to better estimate the

true values of each parameter.

Phycocyanin and acidified chlorophyll-a photopigments and AFDM are used by DEP to

estimate the algae biomass in a stream. This algae biomass, in turn, may be used to

infer characteristics regarding the nutrient status of a stream and also provide

information used to judge the biological condition status of that stream. Phycocyanin is

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a photopigment that, in Pennsylvania, is nearly exclusively descriptive of cyanobacteria.

Chlorophyll-a is a photopigment that is present in all major algal groups.

Subsampling for Phycocyanin

Phycocyanin is subsampled by measuring 30ml of homogenized algae slurry in a 100ml

graduated cylinder then transferring that quantity to a 120ml plastic sample bottle. Swirl

the sample in the 100ml graduated cylinder then pour rapidly into the sample bottle to

help prevent residue from sticking to the graduated cylinder. As with the algae

identification/enumeration subsampling procedure, don’t use de-ionized water to rinse

the graduated cylinder as doing that would introduce error into the final results. A

swirling motion on the 100ml graduated cylinder followed by a quick pour into the

sample bottle will usually prevent sample material from being retained in the graduated

cylinder and will insure that all material is transferred. Sometimes a small quantity of

sample material must be returned to the graduated cylinder from the sample bottle and

a re-pour used to transfer all algae material.

As indicated in Table 2, phycocyanin subsamples require no preservation additives but

does require appropriate low temperature transportation and storage.

The phycocyanin bottle should be labeled with the same GIS Key as was used for the

Algae ID/Enumeration sample, the word “Phycocyanin,” the stream name, and the three

collector-sequence numbers assigned to the subsamples.

Since periphyton algae slurry solution is usually quite concentrated, with regard to

phycocyanin photopigments, BOL will be able to extract three separate phycocyanin

samples from this single 30ml submission – BOL typically does this by re-homogenizing

the 30ml sample and then extracting three ~5ml subsamples. Thus, the three-

subsample replicate requirement is achieved through this single 30ml submission to

BOL.

On the field data sheet, record the three collector-sequence numbers that are to be

used for phycocyanin submissions to BOL. Transfer these collector-sequence numbers,

along with an accurate date and time, the GIS Key, and stream name to the BOL

Microbiology Sample Submission Sheet. Indicate the BOL SAC Code of B031 for the

phycocyanin test on the Sample Submission Sheet. Also, indicate on the BOL

submission sheet that 30ml of sample is supplied and that three 5ml subsamples are

drawn from this single submission (Refer to Appendix A.9 for BOL Microbiology Sample

Submission Sheet).

After subsampling for phycocyanin, place the algae slurry beaker back on the magnetic

stirrer to re-homogenize the sample prior to proceeding to the AFDM subsampling.

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Prepare the bottled phycocyanin sample for transportation as defined in Table 2. If the

biologist hand delivers this sample to BOL, it must be placed into the appropriate

refrigerator in the in the Bacteriological Services Section (consult with lab staff to

confirm the location of this refrigerator). In this case, sample submission sheets should

be hand-delivered to a member of the Bacteriological Services Section so that they are

made aware that a phycocyanin sample has been dropped off – phycocyanin samples

have a short pre-processing holding time (~24 hours, which requires the BOL

microbiologists to preform preliminary processing procedures to stabilize the sample

and protect against degradation of the phycocyanin photopigment. When phycocyanin

samples are delivered by overnight courier, the Bacteriological Services section is made

aware of the arrival of the sample as part of BOL’s normal morning sample check-in

process. Due to the short holding times for pre-processing at BOL, phycocyanin

subsamples should either be hand-delivered at the end of each field day or transported

to BOL by overnight courier at the end of each field day. In either case, phycocyanin

samples should be transported on wet-ice.

Subsampling for AFDM

AFDM is subsampled in a manner very much like that used for acidified chlorophyll-a

(see below). However, preservation procedures differ between the AFDM and acidified

chlorophyll-a subsampling procedures (see Table 2).

AFDM is subsampled by extracting three 2ml replicates of homogenized algae slurry.

This is performed using a 2ml spring loaded Hensen-Stemple Pipette (Wilco part

number 1806-D52) that extracts a subsample from the algae slurry beaker while the

slurry remains on the magnetic stirrer and is continuously homogenized during

subsample extraction.

The contents of the 2ml Hensen-Stemple Pipette is then deposited on a WhatmanTM

GF/F type filter. This is facilitated by a vacuum-draw filtering device (see Figure 5)

which includes a cup-like design that holds the GF/F filter and into which the 2ml

Hensen-Stemple Pipette algae slurry subsample is deposited. While maintaining the

Hensen-Stemple Pipette in an open position, and inside the vacuum-draw filtering

device, rinse the sampler with de-ionized water, also depositing this onto the GF/F

Filter, to be sure that all algae slurry material from the sampler has been deposited on

the filter. After depositing the algae slurry into the filtering device, draw a vacuum on the

filtering device that isn’t greater than 3psi (higher pressure may damage the GF/F filter,

resulting in the loss of periphyton slurry material).

With the vacuum still applied, rinse the side walls of the filter apparatus cup with

deionized water to make sure that all algae material is deposited on the GF/F filter.

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Allow this rinse water to thoroughly drain prior to releasing the vacuum on the filter

apparatus.

Remove the WhatmanTM GF/F filter from the filter apparatus with forceps in a manner

that doesn’t touch or disturb the algae material deposited on the filter. Using two

forceps, one in each hand, carefully fold the round GF/F filter in half so that the algae

material is “sandwiched” inside this now folded half-circle. Continuing to use forceps,

perform one more fold, creating a quartered circle (see Figure 6). Place this folded filter

on a small piece of aluminum foil. Fold the aluminum foil over the filter to contain the

filter entirely within the foil while allowing ample overlap of the foil. Fold the foil edges

over several times to create a sealed envelope around the filter. Each filter replicate is

to be folded into its own foil envelope. Place this foil envelop into a small plastic Ziploc®

bag. Each AFDM filter is to be placed into its own small plastic Ziploc® bag. Place the

three plastic bags into a small paper envelop and seal this paper envelope. This paper

envelope should be labeled with the same GIS Key as was used for the Algae

ID/Enumeration sample, the word “AFDM,” the stream name, and the three collector-

sequence numbers assigned to the subsamples. This envelope is then sealed inside a

Ziploc® plastic bag - multiple paper envelopes may be sealed into a single Ziploc® plastic

bag. Record the three collector-sequence numbers for this sub on the field data sheet.

Transfer the three collector-sequence numbers, along with date and time, the GIS Key,

and stream name to the BOL Microbiology Sample Submission Sheet (Appendix A.9).

The BOL SAC Code for AFDM-a is B027.

Prepare the AFDM sample for transportation as defined in Table 3. When delivering this

sample to BOL, it must be placed into the appropriate -80°C freezer in the

Bacteriological Services Section (consult with lab staff to understand the location of this

freezer). Sample submission sheets should be hand-delivered to a member of the

Bacteriological Services Section so that they are made aware that AFDM samples have

been delivered. AFDM samples may also be delivered to BOL via courier on dry-ice.

However, because of relatively long holding times, AFDM samples may be held for a

few days on dry-ice in coolers and delivered to BOL at the end of the field week.

Quality control measures for AFDM include a laboratory duplicate, a field duplicate, and

a field blank, each done at a rate of once per basin. Field blanks will utilize “ultra-pure

water” instead of the periphyton slurry and will use the same AFDM subsampling

procedures as those described above. Because the term “basin” could cover a broad

geographic area and thus include much of the sampling that occurs in Pennsylvania, for

example the Susquehanna River basin, it will be left to the field biologist to decide the

extent that the term “once per basin” will imply – DEP recommends a conservative

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approach, preferring to submit QA samples regularly (once per week per designated

basin, for example).

Figure 5. Vacuum-draw filtering devices on left and algal slurry beaker on a magnetic

stirrer on the right.

Figure 6. The proper procedure for folding the chlorophyll-a and AFDM filters.

Subsampling for Acidified Chlorophyll-a

Prior to proceeding with the acidified chlorophyll-a subsampling, thoroughly clean the

vacuum-draw filtering cups then rinse with de-ionized water. Acidified Chlorophyll-a is

subsampled in a manner very much like that used for AFDM (see above). However,

preservation procedures differ between the AFDM and acidified chlorophyll-a

subsampling procedures (see Table 2).

Chlorophyll-a is subsampled by extracting three 2ml replicates of homogenized algae

slurry. This is performed using a 2ml spring loaded Hensen-Stemple Pipette (Wildco®

part number 1806-D52) that extracts a subsample from the algae slurry beaker while

the slurry remains on the magnetic stirrer and is continuously homogenized during

subsample extraction.

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The contents of the 2ml Hensen-Stemple Pipette is then deposited on a WhatmanTM

GF/F type filter. This is facilitated by a vacuum-draw filtering device (see Figure 5)

which includes a cup-like design that holds the GF/F filter and into which the 2ml

Hensen-Stemple Pipette algae slurry subsample is deposited. While maintaining the

Hensen-Stemple Pipette in an open position, and inside the vacuum-draw filtering

device, rinse the sampler with de-ionized water, also depositing this onto the GF/F

Filter, to be sure that all algae slurry material from the sampler has been deposited on

the filter. After depositing the algae slurry into the filtering device, draw a vacuum on the

filtering device that isn’t greater than 3psi or persists for longer than 10 minutes, as

greater pressure and/or duration may damage cells resulting the loss of photopigments

(Arar and Collins 1997).

With the vacuum still applied, rinse the side walls of the filter apparatus cup with

deionized water to make sure that all algae material is deposited on the GF/F filter.

Allow this rinse water to thoroughly drain prior to releasing the vacuum on the filter

apparatus.

After the algae slurry water is drawn through the filter, and the algae is deposited on the

GF/F filter, release the vacuum on the filtering device. Fill the filter cup with a small

quantity of de-ionized water so that a few milliliters of water pools on top of the filter.

Into this maintained pool of water, add drop-wise 1ml of saturated MgCO3 solution

preservative. This 1ml of MgCO3 should be uniformly distributed around the pool of de-

ionized water so that it distributes evenly across the entire filter surface. Then re-draw

the vacuum on the filtering device (again, no greater than 3psi) and allow the water and

MgCO3 to drain. New saturated MgCO3 preservative may be obtained from BOL

Microbiological Services at the beginning of the sampling season. This saturated

MgCO3 has a long shelf-life and will remain viable through the entire sampling season

but should be stored in a cool and dry place in the sampling kit.

With the vacuum still applied, rinse the side walls of the filter apparatus cup with enough

de-ionized water to make sure that all MgCO3 is deposited on the GF/F filter. Allow this

rinse water to thoroughly drain prior to releasing the vacuum on the filter apparatus.

Remove the WhatmanTM GF/F filter from the filter apparatus with forceps in a manner

that doesn’t touch or disturb the algae material deposited on the filter. Using two

forceps, one in each hand, carefully fold the circular GF/F filter in half so that the algae

material is “sandwiched” inside the now folded half-circle. Continuing to use forceps,

perform one more fold, creating a quartered circle (see Figure 6). Place this folded filter

on a small piece of aluminum foil. Fold the aluminum foil over the filter to contain the

filter entirely within the foil while allowing ample foil to overlap the filter. Fold the foil

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edges over several times to create a sealed envelope around the filter. Each filter

replicate is to be folded into its own foil envelope. Each acidified chlorophyll-a filter is to

be placed into its own small plastic Ziploc® bag. Place these plastic bags into a single

small paper envelop and seal this paper envelope. This paper envelope should be

labeled with the same GIS Key as was used for the Algae ID/Enumeration sample, the

words “Chl-a Acidified,” the stream name, and the three assigned collector-sequence

numbers. This paper envelope is then sealed inside a Ziploc® plastic bag – multiple

paper envelopes may be sealed into a single Ziploc® plastic bag. Record the three

collector-sequence numbers for this subsample on the field data sheet.

Transfer the three collector-sequence numbers, along with an accurate date and time,

the GIS Key, and stream name to the BOL Microbiology Sample Submission Sheet. The

BOL SAC Code for Chlorophyll-a is B019 – additionally state on the Sample Submission

Sheet that the test is for “Acidified Chlorophyll-a with Pheophytin.” SAC code B019 is

also used for Non-acidified Chlorophyll-a, so care must be taken to clearly communicate

the need for the acidified chlorophyll-a with pheophytin test.

Prepare the acidified chlorophyll-a samples for transportation as defined in Table 3.

When hand-delivering this sample to BOL, it must be placed into the appropriate -80°C

freezer in the Bacteriological Services Section (consult with lab staff to understand the

location of this freezer). Sample submission sheets should be hand-delivered to a

member of the Bacteriological Services Section so that they are made aware that

chlorophyll-a samples have been delivered. Acidified chlorophyll-a samples may be

delivered to BOL via courier on dry-ice. However, because of the holding times involved

(30 days), acidified chlorophyll-a samples may be held for a few days on dry-ice and

delivered to BOL at the end of the field week.

USEPA recommends that quality control measures for chlorophyll-a include a laboratory

duplicate, a field duplicate, and a field blank, each done at a rate of once per basin

(USEPA 2013). Field blanks will utilize “ultra-pure water” instead of the periphyton slurry

and will use the same acidified chlorophyll-a subsampling procedures as those

described above. Because the term “basin” could include a broad geographic area and

thus include much of the sampling that occurs in Pennsylvania, for example the

Susquehanna River basin, it will be left to the field biologist to decide the extent that the

term “once per basin” will imply – DEP recommends a conservative approach,

preferring to submit QA samples regularly (once per week per designated basin, for

example). USEPA suggests an acceptance criteria of 25% relative percent difference

for the laboratory and field duplicates, and 0.00µg/L ±0.11µg/L for the field blank

(USEPA 2013).

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Subsampling for Cyanotoxins

After the acidified chlorophyll-a subsamples are collected, continue homogenizing the

algae slurry on the magnetic stirrer. Pour 250ml of this homogenized slurry into a 500ml

graduated cylinder. Swirl this slurry around the graduated cylinder and quickly pour it

into a Whirl-Pak® bag to prevent algae material from adhering to the sides of the

graduated cylinder. Again, don’t rinse down with additional de-ionized water as that

would dilute the sample and introduce error into the final results. Seal the Whirl-Pak®

bag then transport and store as prescribed in Table 2.

DENSIOMETER AND COMPASS DATA COLLECTION

In smaller streams, the riparian vegetative cover may intercept much of the incident

solar radiation that may otherwise be available for supporting primary production in the

lotic habitat. Trees situated along the riparian corridor are very effective at appreciably

reducing the quantity of light reaching the stream (Hill and Roberts 2009). There is great

potential for a combination of both light and nutrients to colimit algal growth (Hill 1996).

Additionally, the East/West or North/South orientation of the stream reach being

sampled will also seasonally contribute to the quantity of incident sunlight on the stream

reach. Consequentially, a measurement of canopy closure (thus inferring intercepted

solar radiation) over the stream reach, and stream orientation, should be made by the

biologist and considered, in conjunction with water chemistry measures, when analyzing

the results of algae sampling.

Stream orientation is measured with a compass. Facing downstream at the

Densiometer “1/2 Downstream” location (refer to either the QBE or QMH Field Data

Sheets in Appendices A.6 and A.7), site the compass along the thalweg in the

downstream direction and orient the compass with magnetic north. Record this sighted

azimuth direction clockwise from magnetic north (in degrees) on the data sheet.

Stream canopy closure is measured with a Concave Spherical Densiometer (for

example, part number 43888 available from Forestry Suppliers, Inc at www.forestry-

suppliers.com). Procedures for estimating canopy closure follow Platts et al. (1987).

Measuring canopy closure is suggested over measurement of canopy density since

canopy closure measures are less affected by seasonality (Fitzpatrick et al. 1998).

Vegetative canopy closure is the area of the sky bracketed by vegetation. The

densiometer's concave mirror surface has 37 grid intersections forming 24 squares. To

eliminate bias from overlap, only 17 of the 37 grid intersections are used as recording

points. The 17 grid intersections to be used are delimited by taping a right angle on the

densiometer (see Figure 6).

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For streams measuring less than 25 meters across (82 feet), densiometer readings are

recorded at four locations along the mid-reach transect – these being, facing the right-

descending bank, facing mid-channel downstream, facing mid-channel upstream, and

facing left-descending bank. This is accomplished, for example at the right-descending

bank, by facing the right wetted edge of the stream and holding the densiometer about 1

foot from the shoreline and about 1 foot above the water surface (see Figure 7). With

the leveled (bubble level) densiometer now pointed toward the bank (taped right-angle

points toward the recorder) and with the observer’s head reflection near the top grid line

(see Figure 6) then determining canopy closure at that location. Thereafter, canopy

closure is counted at each of the remaining mid-reach transect locations in a similar

fashion. Canopy closure count is equal to the number of densiometer line intersections

(maximum of 17) that are surrounded by vegetation (representing canopy closure) or

intercepted. Finally, the count for each location is then summed together and divided by

68 (4 x 17) to calculate a percent closure. For example, if the canopy closure count at

the right-descending bank is 15, the count at mid-channel facing upstream is 3, the

count at mid-channel facing downstream is 5, and the left-descending bank count is 16,

then the canopy closure value for the site is determined by the following: [(15 + 3 + 5 +

16)/68] x 100 = 57% (round to nearest whole percent).

For streams larger than 25 meters across the same procedure is used except eight

recordings are made. In addition to right bank, left bank, and mid channel (upstream

and downstream) locations, mid-reach upstream and downstream densiometer

measurements are additionally made at the ¼ and ¾ intervals on the mid-reach

transect. Canopy closure or density scores are summed and divided by 136 (8 x 17) to

calculate a percent closure.

Figure 6. Concave spherical densiometer with placement of head reflection, and 17

observation points. From Platts et al. (1987).

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Figure 7. Proper use of the densitometer at the right descending bank location.

LABORATORY PROCEDURES

Although strict adherence to the field data collection procedures is necessary to achieve

comparability between samples, field data collection represents only a portion of that

which is needed to insure robust periphyton data results. Laboratory procedures are

also extremely important and must also be defined and strictly adhered to if sample

comparability is to be achieved. The purpose of this section is to define those laboratory

procedures needed to process periphyton related samples. These laboratory

procedures include those applied by BOL as well as those that must be applied by

contracted labs. These protocols are as follows:

• Phycocyanin by Fluorometry

• Acidified Chlorophyll-a by Fluorometry with Pheophytin

• Ash-Free Dry Mass by SM10200-I

• Algae Identification and Enumeration

• Quality Assurance (QA) Audit of Algae Identification and Enumeration.

At present, DEP’s Algae Monitoring Program processes Phycocyanin, Acidified

Chlorophyll-a (which includes Pheophytin reporting), and Ash-Free Dry Mass samples

at its BOL facility. The laboratory procedures used to process these samples are

extensively defined in BOL documentation and may be obtained from that DEP bureau.

BOL methods currently used include:

• Phycocyanin by Fluorometry Rev 001, effective November 2017

• [Acidified] Chlorophyll-a in NPW by SM10200H 1 & 3 (Fluorometric Method Rev

002, effective 3/27/2017

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• Ash-Free Dry Mass by SM10200-I Rev 001, effective 2/26/2016

• Plankton Identification and Enumeration Rev 002, effective 5/7/2015.

Algae Identification and Enumeration samples collected by DEP are processed through

private companies, academic institutions, as well as through DEP BOL. Because these

identification samples are processed through several different facilities the reader is

directed to both the BOL Plankton Identification and Enumeration document as well as

to the Algae identification and Enumeration Protocol description provided below.

Further, the use of multiple identification and enumeration processing facilities obligates

DEP to employ a rigorous QA Audit process designed to insure consistency, and thus

comparability, between the results produced by the various labs. This QA Audit process

is also discussed below.

General Description of Algae Identification and Enumeration Procedure

The identification and enumeration of periphyton samples collected by either the QMH

or the QBE periphyton field procedures requires that two separate procedures be

performed to completely characterize the sample. These include a Palmer-Maloney

Chamber procedure as well as a Diatom Slide procedure.

The Palmer-Maloney Chamber procedure, sometimes referred to by lab technicians as

the “wet count,” or “soft algae count,” identifies and counts all algae taxa to the genera

level, at a minimum, at moderate magnification (e.g., 100x or 400x). However, if certain

morphological structures are observable during this procedure, identifications may be

made to the species, or sometimes variety, taxonomic level. It is important to recognize

that during the Palmer-Maloney procedure, diatom counts are also made. Diatoms,

however, are only characterized as living or as dead and are not taxonomically

identified in this procedure. Palmer-Maloney Chambers used in this process must be

calibrated.

As previously mentioned, the Palmer-Maloney procedure is sometimes referred to as

the “wet count” or the “soft algae count” as a way of distinguishing it from the Diatom

Slide count. The “wet count” name originates from the fact that the submitted

identification subsample is further subsampled by placing a small homogenized quantity

of liquid (hence the term “wet”) into a Palmer-Maloney chamber for microscopic

inspection. The term soft algae is also often applied to this procedure because the algae

taxa identified and counted in this procedure frequently contain those possessing only

soft cell tissues that would otherwise be destroyed as part of the nitric acid digestion

used to create slides for the diatom slide procedure. Examples of such “soft algae”

commonly identified in samples collected from Pennsylvania streams and rivers include

cyanobacteria or green algae taxa. However, Palmer-Maloney results do sometimes

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include other “soft algae” taxa, which are less commonly found in Pennsylvanian

freshwater periphyton samples.

In contrast to the Palmer-Maloney procedure, diatoms are identified to the species, and

often to variety, taxonomic level in the Diatom Slide procedure. In this procedure, liquid

from the subsample is processed with nitric acid to dissolve away all soft tissues in the

sample thereby exposing the glass frustules of the diatoms. A detailed microscopic

analysis of the diatom glass frustule morphology is then performed to make the species

or variety identification at high magnification (1000x oil immersion). The identifications

and counts obtained during this procedure produce what is often referred to as the

“permanent slide” result. Permanent slides are a product of the procedure referred to as

the Preparation of Diatom Slides Using NaphraxTM Mounting Medium, as listed below.

DEP requires the identification laboratory to produce three permanent slides per

submitted algae identification subsample. These three permanent slides are maintained

by DEP as voucher specimens and are also used in the QA audit.

As a separate option, the BOL Algae Identification and Enumeration procedure may be

used to produce genera level taxonomic identifications and enumerations. Also, the

BOL procedure only provides results corresponding to the Palmer-Maloney procedure

described above. These results, though less informative than those produced by the

complete procedure, which combines results from both the Palmer-Maloney and Diatom

Slide procedures, are used when an algae community result is needed from a

laboratory in a very short period of time (historically, DEP has experienced that

complete sample results from laboratories are often obtained only after many months of

processing time at those facilities). If this BOL option is to be used, modifications to the

standard BOL Algae Identification and Enumeration procedure are needed. These

modifications include counts of living and dead diatoms during the Palmer-Maloney

count phase and the fact that the BOL method specifies “phytoplankton… samples”

whereas a periphyton sample is submitted. [Note: living diatoms are those that are

observed to contain soft tissue within the frustule during the Palmer-Maloney procedure

whereas those that contain no such soft tissue are considered to be dead at the time of

sampling. Also, the QA procedures expected of Participating Labs (see below) must be

required of samples processed by DEP BOL.]

Algae Identifications that use both the Palmer-Maloney soft algae analysis as well as

the diatom slide analysis will rely, in general, upon the Protocols for the Analysis of

Algal Samples Collected as Part of the U.S. Geological Survey National Water-Quality

Assessment Program, Report No. 02-06 by Charles et al. 2002 (available at

https://water.usgs.gov/nawqa/protocols/algprotocol/algprotocol.pdf). The protocols from

Report No 02-06 that are routinely applied, in part or in whole, by DEP include:

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• P-13-58: Tracking of Algae Sample Analysis – Figure 1 (Sample process and

analytical flow)

• P-13-48: Subsampling Procedures for USGS NAWQA Program Periphyton

Samples

• P-13-42: Diatom Cleaning by Nitric Acid Digestion with Microwave Apparatus

• P-13-49: Preparation of Diatom Slides Using NaphraxTM Mounting Medium

• P-13-50: Preparation of Algal Samples for Analysis Using Palmer-Maloney Cells

• P-13-39: Analysis of Diatoms on Microscope Slides Prepared from USGS

NAWQA Program Algae Samples

• P-13-63: Analysis of Soft Algae and Enumeration of Total Number of Diatoms in

USGS NAWQA Program Quantitative Targeted-Habitat (RTH and DTH)

Samples.

The rationale for performing the QBE and QMH Periphyton Sampling Procedures as

well as the Algae Identification and Enumeration Procedures, as based upon the USGS

NAWQA protocols, is to produce results of the quality needed for the development of a

trophic index, that needed for a diatom based Multi-Metric Index (MMI), as well as that

needed to provide information that contributes to aquatic life use considerations. These

procedures provide community-level data as well as the identification of pollution

sensitive or insensitive diatoms. To achieve segregation between potential levels of

pollution tolerance within the diatom group, identification to the species or variety

taxonomic level is required in the Diatom Slide Procedure. In short, diatoms, when

identified to species or a variety of taxonomic levels, provide information concerning the

nutrient and organic enrichment condition of natural waters (Porter et al. 2008) and will

also serve as indicators for determining biological condition gradients and developing

nutrient criteria (Hausmann et al. 2016).

Detailed Description of Analytic Services for Algae Identification and

Enumeration

DEP requires that algae identification and enumeration of benthic algae samples be

done in a way that produces results “comparable” to those provided by The Academy of

Natural Sciences (ANS). ANS produces results as described by the previously

mentioned protocols for the analysis of algal samples collected as part of the U.S.

Geological Survey National Water-Quality Assessment Program (NAWQA Protocol,

Charles et al. 2002).

The word “comparable,” in the above paragraph, is emphasized because DEP is

primarily interested that the results produced by the identification and enumeration

process completely harmonize with those produced by the ANS. Consequentially, there

may be a limited opportunity for flexibility regarding the exact procedures by which the

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identification and enumeration laboratory results are produced. This creates the

possibility for the processing laboratory to slightly modify the procedures of performing

identification and enumeration as detailed by the NAWQA Protocol so long as that

laboratory does so in close consultation with, and approval by, DEP.

Algae identification and enumeration results will be achieved by performing the following

process steps. It is important to note that this process requires a close collaboration

between DEP and the Participating Laboratory (PL). To facilitate this collaboration, the

areas of responsibility are indicated in each step. Also, the applicable NAWQA protocol

is also indicated, where appropriate. The process steps will include:

1. Collection of periphyton sample and division into 100ml subsample bottles

preserved with 5% to 10% formaldehyde. Service provided by DEP as

described above.

2. Preparation of permanent diatom microscope slides via acid digestion to

produce “clean” identification and enumeration slides as well as three

permanent archive (voucher) slides for each sample – refer to NAWQA

Protocols P-13-58 Figure 1, P-13-48, P-13-42, and P-13-49. In general, a

permanent slide diatom specimen density should be constructed so that

between 5 and 10 diatom specimens are observable in a single microscope

field at 1000x magnification (recommendation by Charles et al. 2002).

3. 300 Natural Unit ID/Count Analysis of soft algae and diatoms in a calibrated

Palmer-Maloney chamber for each sample – refer to NAWQA Protocol P-13-

58 Figure 1, P-13-48, P-13-50 and P-13-63. Identification of soft algae to

lowest possible taxonomic level (at discretion of analyst but minimum of at

least general level expected). Differentiate diatoms into living (containing

cytoplasm) and dead (containing no cytoplasm). Identifications and counts

must be harmonized. Identifications and counts provided by the PL.

Harmonization coordinated between DEP and PL as defined in the

harmonization section below.

4. 600 Valve ID/Count Analysis of diatom microscope slides for each sample –

refer to NAWQA Protocol P-13-39. Identification to species or variety

taxonomic level. Identifications and names must be harmonized.

Identifications and counts provided by the PL. Details on coordinating

harmonization between DEP and PL is defined in the harmonization section

below.

5. Blind Duplicate and QA Audit. DEP will provide blind duplicates samples to

the PL. The QA Audit process for these blind duplicates is defined below.

Also included in this QA process is the collecting of taxa specific

photomicrographs by the PL, as defined in the QA Audit Process below. The

PL will also complete its own internal QA Audit process to help insure its own

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method quality, particularly as it relates to the processing of the Palmer-

Maloney slides and the diatom slides. The PL will design and implement its

internal QA Audit process in consultation with DEP. The PL will share the

results of their internal QA Audit process with DEP.

6. Reporting Identification and Enumeration Results to DEP. The PL will make a

final identification and enumeration results report (preliminary reports are

made prior to the completion of the QA Audit process) to DEP in the format

discussed below. The final results report will include all necessary changes to

preliminary results reports provided by the PL as required by the outcome of

the Blind Duplicate and QA Audit processes. This is a collaborative process

between the PL and DEP.

Blind Duplicates and Other QA Audit Process:

Blind Duplicate QA Audit processing success will be determined with reference to

published and peer reviewed scientific literature. The scientific literature referenced and

currently consulted by DEP for this QA Audit process is included in the Literature Cited

section of this document and is summarized below. However, the rapid pace at which

the algae science is progressing will require DEP and the PL to frequently review and

discuss the current body of applicable scientific literature and accepted procedures.

Both the PL and DEP will be expected to make on-going adjustments to incorporate the

recent changes and advances in the applicable scientific literature. Scientific literature

for performing the QA Audit process includes:

• Barbour et al. 1999

• Kahlert et al. 2009

• Kelly 2001

• Whittaker and Fairbanks 1958

• Wolda 1981

• PCER 1998

The body of scientific literature consulted suggested that the appropriate procedure for

determining the quality of reported diatom identifications and enumerations is to use

indices of similarity to compare the results of a sample with those from a corresponding

blind duplicate sample, also processed by the PL, as well as a comparison with results

independently produced by an auditor. Whereas the blind duplicate is intended to

demonstrate the ability of the PL to replicate their own results, the comparison to the

auditor is intended to confirm the accuracy of identification and enumerations. Deciding

who in DEP or what outside laboratory will serve as the auditor will be decided by DEP

at the beginning of each project. DEP typically submits blind duplicates at a rate of 10%

of the regular sample load. If blind duplicate or auditor similarity comparisons fail to

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pass (see below for pass scores), DEP must consult with the PL to resolve this quality

issue before incorporating data into the DEP Algae Database.

The QA process will also require the PL to provide photomicrographs (in a digital format

agreed upon by both the PL and DEP) to DEP at the time of data submission. These

photomicrographs will include an image example of each taxa type identified in the DEP

sample set and must be photographed from the DEP sample set. Diatom identification

images must be photographed at 1000x and must clearly show all structures needed to

confirm the identification to the appropriate species or taxonomic level. Images collected

during the Palmer-Maloney 300 Natural Unit process must be photographed at the

appropriate magnification needed to make the identification and must also clearly show

all structures needed to confirm the identification. Photomicrographs must include a

calibrated scale bar. Photomicrograph files must be entitled to include the taxon name,

magnification, date of photograph, and DEP sample from which it was obtained. Again,

the PL is required to provide only one type image for each taxon identified in the entirety

of the DEP sample set (per contract or Purchase Order), regardless of how many times

the taxa is re-identified throughout the sample set.

The indices of similarity that DEP uses in the QA Audit process will include the

following:

• Percent Community Similarity of Diatoms (PS) – this is also referred to as the

Whittaker and Fairbanks (1958) similarity and is recommended by USEPA

RBP. The USEPA RBP suggests a percent similarity score of > 75% to pass.

This index is based upon relative abundance and places more emphasis

upon the more abundant taxa while de-emphasizing the rare taxa. Computed

as:

PS = 100 – ( 0.5Σ│ai – bi │ )

where:

ai = percent abundance of species i in sample A

bi = percent abundance of species i in sample B.

• Bray-Curtis Similarity Index (D) – This index is also based upon relative

abundance. Note that Whitaker-Fairbanks Percent Community Similarity and

the Bray-Curtis Similarity Index are numerically equivalent. The scientific

literature (Kelly 2001) suggests that a Bray-Curtis score of > 60% is generally

considered to be a good agreement between the taxonomist and the auditor,

especially when taxa diversity is relatively high. However, this pass score

should be increased to 70% when taxa diversity is relatively low. Computed

as:

3-130

D = Σ min ( ai , bi )

where:

ai = percent abundance of species i in sample A

bi = percent abundance of species i in sample B.

Harmonization:

The harmonization of algae sample identification and enumeration procedures between

DEP and the PL, and between processing laboratories, is essential to the success of

DEP’s algae monitoring and method development effort. DEP has experienced

inconsistencies between laboratories regarding their reported identifications and counts.

Such inconsistencies were also noted by USEPA during the analysis of their 2008/2009

National Rivers and Streams Assessment (NRSA) diatom data (Lee et al. 2017). There

are several possible reasons for these inconsistencies, many of which are described by

Kahlert et al. (2009). Kahlert et al. (2009) suggests that harmonization is likely

imperative to achieve consistency between laboratories and taxonomists. Kahlert et al.

(2016) suggests methods by which this is may be achieved. Currently, DEP recognizes

that to achieve proper harmonization, the following operational considerations and

activities must be continuously reviewed and coordinated between both DEP and the

PL:

• The body of current taxonomic references and literature to which the

taxonomist refers to properly identify algae taxa

• Defining appropriate species complexes into which are grouped problematic

identification OTU’s (Operational Taxonomic Units) (Refer to Lee et al. 2017)

as well as summarizing taxa that may easily be misidentified (Refer to Kahlert

et al 2016).

• All laboratory procedures involved with the processes of subsampling, and

identifying/enumerating algae samples

• Possible use of taxonomic intercalibration exercises and other harmonization

procedures (Kahlert et al. 2009, Kahlert et al. 2016).

Because of the importance of diatom related data to DEP, the proper harmonization of

diatom taxa is especially emphasized. In demonstration of the importance of

harmonization, refer to Kahlert et al 2009 in which the authors suggest that proper

diatom taxa harmonization, rather than long experience with diatoms, is more important

for achieving similarity between samples (and between taxonomist and auditor).

Initial steps for achieving harmonization between DEP and the PL will involve the

following at the beginning of the project:

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• DEP will provide to the PL the currently acceptable taxa list. This taxa list will

include reference codes for each taxa along with authorship information.

• DEP and the PL will review the PL’s identification and enumeration procedures.

Notes regarding taxa lists, DEP naming conventions, and identifications:

• The taxa list to which DEP refers, in general, is typically that offered by The

Academy of Natural Sciences in their current list of Diatom and Non-Diatom (soft

algae) This current downloadable Academy taxa list is available at

http://diatom.ansp.org/Taxa.aspx. DEP also frequently refers to Diatoms of the

United States (DOTUS) for taxa nomenclatural details (available at

https://westerndiatoms.colorado.edu/). The Academy list is preferred because it

is downloadable and is easily incorporated into a database.

• By DEP naming conventions, taxa names must include the source of authorship

as part of the name. For example, Achnanthidium minutissimum (Kützing)

Czarnecki, Achnanthidium minutissimum var. gracillima (Meister) Bukhtiyarova,

or Phormidium autumnale (Agardh) Trevisan ex Gomont. This convention is used

by The Academy and must be rigidly adhered to by the PL, thus facilitating

compatibility with the DEP database.

• For identifications, DEP also consults DOTUS (available at

https://westerndiatoms.colorado.edu/), AlgaeBase (available at

http://www.algaebase.org/), the Diatom New Taxon File maintained by The

Academy of Natural Sciences (available at http://dh.ansp.org/dntf), the Catalogue

of Diatom Names maintained by the California Academy of Sciences (available at

http://researcharchive.calacademy.org/research/diatoms/names/index.asp ). And

USGS Biodata (available at https://aquatic.biodata.usgs.gov).

• The above notes, and institutional references, are not exhaustive and may

include others when deemed appropriate.

The above represents initial harmonization steps that must be performed at the

beginning of each project. Bear in mind that changes in algae taxonomy occur

frequently and issues with problematic identifications will always exist. Consequentially,

DEP and the PL will continuously confer so that they together may contend with

taxonomy changes and problematic identifications as frequently as is deemed

necessary by either of the two parties.

DEP will appoint an Auditing Laboratory at the beginning of each project. DEP will

advise the PL as to the identity of the Auditing Laboratory at the beginning of the

project.

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Algae Identification and Enumeration Results Reporting, Data Formatting, and PL

Internal Diatom and Palmer-Maloney Slide Processing QA Audit Report

Requirements:

DEP will require that certain types of data be generated by the PL when processing

Algae ID and enumeration samples. Data transfer between DEP and the PL must follow

a specific format, as demonstrated by the tables presented in this section. This specific

table format is necessary to efficiently support data transfer into DEP’s Microsoft®

Access based Algae Database. Three data table transfers are needed to properly

exchange data between DEP and the PL. These tables include:

• Algae Identification and Enumeration Laboratory Submission Data Sheet,

provided by DEP to the PL upon submission of samples

• Palmer-Maloney and Diatom Slide Applied Factors Table, provided by the PL to

DEP

• Identification/Enumeration Results Table, provided by the PL to DEP.

Results will not be fully incorporated into the DEP Algae Database until all PL

processing quality issues (as per blind duplicate results and auditor comparison results)

are resolved and corrected data is received from the PL.

Each of the Results Reporting Tables will be defined and formatted as follows:

1. Algae Identification and Enumeration Laboratory Submission Data Table. DEP

will transfer samples and the Algae Identification and Enumeration Laboratory

Submission Data Table to the PL at the beginning of the laboratory processing

phase of the project. This data table will exist in the form of a Microsoft® Access

data table (or Excel data table if necessary). Refer to Appendix A-8 for the format

of this table. The following fields are required in this table to fully describe each

sample to the PL (origin of information from the DEP Algae Database are

included in parenthesis):

• DEP Algae Sample GIS Key ID. Access Foreign Key that references the

corresponding Primary Access Key in the DEP Algae Database Samples

Table. Configuration of this GIS KEY ID is YYYYMMDD-HHMM-Collector.

For example, Algae Sample GIS Key ID’s have, in the past, included

20140813-0800-jbutt, and 20140821-0800-ggocek.

• Date sample was collected. (Duplication from the DEP Algae Database

Samples Table).

• Sample Type. QMH (Qualitative Multi-Habitat) or QBE (Quantitative

Benthic Epilithic) periphyton sample.

• DEP Algae Sampling Site ID number. Access Foreign Key that references

the corresponding Primary Access Key in the DEP Algae Database Sites

Table

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• Stream Name. (Duplication from the DEP Algae Database Sites table)

• Location Description. (Duplication from the DEP Algae Database Sites

Table).

• County. (Duplication from the DEP Algae Database Sites Table)

• Latitude and Longitude. (Duplication from the DEP Algae Database Sites

Table).

• Benthic Surface Area (cm2). Applies only to QBE Periphyton samples

(Duplication from the DEP Algae Database Samples Table).

• Algae Slurry Total Sample Volume (ml). Applies only to QBE Periphyton

samples (Duplication from the DEP Algae Database Samples Table).

• ID Bottle SubSample Volume (ml). (Duplication from the DEP Algae

Database Samples Table). Typically 100ml.

• Formaldehyde Added to ID SubSample Bottle (ml). (Duplication from the

DEP Algae Database Samples Table).

• Note: Transfer of all this data to the PL is necessary. Location of sample

may help the algae taxonomist in deciding species or variety level taxa ID

due to possible geographic floristic variations. Surface areas and volumes

are required by the PL to perform sample specific calculations that effect

values of the Palmer-Maloney Correction Factors and the Diatom Slide

Applied Factors.

2. Palmer-Maloney and Diatom Slide Applied Factors Table. The PL will transfer to

DEP sample specific values as determined during the analysis of the Palmer-

Maloney slide and reported in the Palmer-Maloney and Diatom Slide Applied

Factors Table (as a reminder, any Palmer-Maloney Chambers used to produce

for DEP must be calibrated). To facilitate this data transfer process, DEP will

provide a Microsoft® Access data table (or Excel data table if necessary)

template to the PL at the beginning of the identification/enumeration project,

which the PL will then populate and return to DEP. An abbreviated example of

this table is provided in Table 3 and demonstrates real data provided by The

Academy of Natural Sciences for samples collected in 2013.. This table will

include the following fields to fully characterize each sample to DEP:

• DEP Algae Sample GIS Key ID. Access Foreign Key that references


Table. Obtained by PL from the Algae Identification and Enumeration

Laboratory Submission Data Table

• Field Dilution Correction Factor (FDCF). DEP submits to the PL a 100ml

algae ID/enumeration subsample bottle to which is added a formaldehyde

preservative. This correction factor is determined by the PL from values

provided on the Algae Identification and Enumeration Laboratory

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Submission Data Table and is computed as: (Volume of ID/enumeration

subsample (ml) + Volume of Formaldehyde added (ml)) / Volume of

ID/enumeration subsample (ml). For example, if volume of ID/enumeration

subsample = 100ml and the amount of formaldehyde added to that

subsample is 5 ml, then this correction factor = (100ml + 5ml)/100ml =

1.05.

• Subsample Dilution Correction Factor (SDCF). If the PL needs to make a

dilution to the submitted ID/enumeration subsample bottle the SDCF is

recorded in this field by the PL. SDCF is computed as: (Volume of original

subsample ID/enumeration bottle (ml) + Volume of distilled water added

(ml)) / Volume of original subsample ID/enumeration bottle(ml)).

• Palmer-Maloney Dilution Correction Factor (PMDCF). If the PL needs to

make a dilution during the Palmer-Maloney analysis, the PMDCF will be

recorded in this field by the PL. Computation is similar to those field

dilution correction factors demonstrated above.

• Palmer-Maloney Microscope Field of View Volume, and Palmer-Maloney

Number of Fields Observed. Each field of view in the microscope will

contain a certain volume, and is dependent upon the Palmer-Maloney

chamber depth and the diameter of the field of view of the microscope. For

example, from DEP sample 20130424-0800-jbutt, the PL microscope’s

field of view volume in the Palmer-Maloney was 0.000048497ml/field of

view. The number of fields of view is a record of the number of fields of

view that must be completely counted to reach, or slightly surpassed, the

300-natural units threshold required by the Palmer-Maloney analysis.

• Palmer-Maloney Total Volume of all Fields Observed. This value is

calculated by the PL by multiplying the field of view volume by the quantity

of fields observed. For example, from the DEP sample 20130424-0800-

jbutt, this value was (0.000048497ml/field of view) x (10 fields of view) =

0.0004897ml.

• Algal Slurry Total Sample Volume (ml). Duplicated from the Algae

Identification and Enumeration Laboratory Submission Data Sheet.

• Benthic Surface Area of sample (cm2). Applies only to QBE Periphyton

samples Duplicated from the Algae Identification and Enumeration

Laboratory Submission Data Sheet.

• Cell Density Factor (CDF). This factor is computed by CDF = (FDCF *

SDCF * PMDCF * Algae Slurry Total Sample Volume) / (Total Volume of

all Fields observed in the Palmer-Maloney * Benthic Area Sampled).

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Table 3. Example of the Palmer-Maloney and Diatom Slide Applied Factors Table for

2013 DEP algae Samples processed by The Academy of Natural Sciences.

3. Identification/Enumeration Results Table. The PL will transfer to DEP, taxa

specific values for each sample as determined during the analysis of the Palmer-

Maloney chamber and diatom slides and reported in the

Identification/Enumeration Results Table. To facilitate this data transfer process,

DEP will provide a Microsoft® Access data table (or Excel data table if

necessary) template to the PL at the beginning of the identification/enumeration

project, which the PL will then populate and return to DEP. An abbreviated

example (shows only the results from a single sample and not those generated

from all samples) of this table is provided in Table 4. This table will include the

following fields to fully characterize each sample to DEP:

• DEP Algae Sample GIS Key ID. Access Foreign Key that references


Table.

• Taxonomic NADED ID Number. Access Foreign Key that references

corresponding Primary Key in the DEP Algae Database Taxa List Table.

North American Diatom Ecological Database (NADED) ID numbers,

unique for each taxa name-taxa author, are available from The Academy

of Natural Science current taxa list at http://diatom.ansp.org/Taxa.aspx

and are also found in the Diatoms of the United States website available

at https://westerndiatoms.colorado.edu/ . For example, Achnanthidium

minutissimum (Kützing) Czarnecki is listed as NADED 1010,

Achnanthidium minutissimum var. gracillima (Meister) Bukhtiyarova as

NADED 1063, and Phormidium autumnale (Agardh) Trevisan ex Gomont

as NADED 890028 in the Academy current taxa list.

• Algae Taxon Name with Author Reference. Refer to the Harmonization

section, above, for requirements.

• Natural Units Counted in Palmer-Maloney. Natural Units are counted for

each sample during the soft algae analysis in a Palmer-Maloney counting

20130424-1100-jbutt 700 185.4 1.05 1 00 2.00 0 000048497 10 0.00048497 16349.1

20130424-1400-jbutt 700 185.4 1.05 1 00 2.00 0 000048497 13 0.00063046 12576 2

20130424-0800-jbutt 700 185.4 1.05 1 00 1.00 0 000048497 10 0.00048497 8174 5

20130916-0800-jbutt 1784 556.2 1.05 1 00 2.00 0 000048497 43 0.00208537 3230 0

20130916-1400-jbutt 1099 556.2 1.05 1 00 1.00 0 000048497 47 0.00227936 910.2

20130916-1100-jbutt 1109 556.2 1.05 1 00 1.00 0 000048497 66 0.00320080 654.1

20130917-1100-jbutt 1214 556.2 1.05 1 00 1.00 0 000048497 47 0.00227936 1005 5

20130917-1400-jbutt 1834 556.2 1.05 1 00 4.00 0 000048497 37 0.00179439 7717 9

20130917-0800-jbutt 1014 556.2 1.05 1 00 2.00 0 000048497 39 0.00189138 2024 2

PADEP Algae Sample

GIS Key ID (PADEP)

FDCF -

Field

Dilution

Correction

Factor

SDCF -

SubSample

Dillution

Correction

Factor

Algae Slurrry

Total Sample

Volume (ml)

Benthic

Area

Sampled

(cm2)

CDF - Cell

Density

Factor in

Palmer-

Maloney

PMDCF -

Palmer-

Maloney

Dilution

Correction

Factor

Microscope

Field of View

Volume in

Palmer-

Maloney (ml)

Number of

Fields

Observed

in Palmer-

Maloney

Total Volume

of all Fields

Observed in

Palmer-

Maloney

3-136

chamber. Natural Units are natural groupings of algae such as an

individual filament, colony, or isolated cell and represent the usual

grouping of cells that occur in a natural setting. Each such natural

grouping is counted as one natural unit. For example, a colony of

microcystis, which may contain dozens or perhaps hundreds of cells, is

counted in a soft algae sample as one natural unit. This counting

procedure is used for all non-diatom taxa. Living and dead diatoms,

regardless of their colonial grouping, are counted so that each cell is a

natural unit, even when they are observed in colony form. Living diatoms

must be distinguished from dead diatoms. This unit exists to prevent

colonies or filaments from dominating the result of the 300 Natural Unit

ID/count, thereby enabling a good representation of the taxa diversity in

the soft algae sample.

• Number of Cells Counted in Palmer-Maloney. The number of cells are

counted during the soft algae analysis in a Palmer-Maloney counting

chamber. In soft algae samples, an attempt to count (or estimate, if a

count of a colony is impractical) the number of cells will be performed.

Each diatom frustule or valve is counted as a cell in a soft algae sample.

Care needs to be exercised in producing this result, especially when

colony estimation is performed since the cell count is used to derive cell

abundance and biovolume in the permanent slide results.

• Number of Valves Counted in Diatom Slide for each Taxa. These are the

number of diatom valves counted for each taxa in the microscope fields of

view during the analysis of the diatom slide. Diatoms contained on these

slides have been treated to remove soft tissues from the sample, thereby

rendering the silicate structure of the diatom frustules highly visible in a

microscopic study.

• Total Number Valves Counted on Diatom Slide. Equal to the sum of all

valves for each taxa counted in the microscope fields of view during the

analysis of the diatom slide. Microscope fields of view on the diatom slide

are completely counted until a threshold quantity of 600 total valves is

reached or slightly surpassed.

• Diatom Valve Relative Abundance. This computed value is equal to the

number of diatom valves counted in each taxa on the diatom slide divided

by total number of valves counted on the diatom slide.

• Number of Living Diatoms Counted in Palmer-Maloney Analysis. This is

the quantity of living diatom cells counted as part of the Palmer-Maloney

identification and Enumeration Process. This value is derived for the entire

sample and consequentially is the same for each taxa in the sample. [Special Note: At this point it is necessary to describe, in detail, the difference between

living and dead diatoms. By definition, Undifferentiated Diatoms - Living (NADED

3-137

249999) are the number of diatoms containing cytoplasm counted during the Palmer-

Maloney analysis. Because of the presence of cytoplasm, these diatoms are to be

considered as having been alive at the time the sample was collected in the field. Note

that the cytoplasm in a living cell, along with all the cell’s organelles, may be reduced into

a small packet, of sorts, due to the formaldehyde preservation and/or the storage time.

Whereas, by definition, Undifferentiated Diatoms – Dead (NADED 249998) are the

number of diatoms or diatom valves counted during the Palmer-Maloney analysis that

don’t contain any evidence of cytoplasm. Because these diatoms or valves do not contain

cytoplasm these diatoms are to be considered as having been dead at the time the

sample was collected.] Living and dead diatoms are identified and counted as

taxa types in the Palmer-Maloney Identification and Enumeration process.

Because living and dead diatoms contribute to the Natural Unit count, this

distinction must be made to correct the diatom slide count to eliminate that

proportion of diatoms presumed to have been dead at the time the sample

was collected in the field (in other words, the diatom slide result must be

adjusted to estimate only the quantity of diatoms that were alive at the

time of sampling). This data field is needed because the nitric acid

treatment used to prepare the diatom slide dissolves all soft tissues

making it impossible to distinguish between diatoms that were living

versus those that were dead at the time the sample was collected.

Consequentially, the taxonomist must ID and count all observed diatoms

then make the correction discussed in the Proportioned Diatoms field.

• Proportioned Living Diatom Valves. This computed value is equal to

Diatom Valve Relative Abundance value multiplied by the number of

Undifferentiated Diatoms – Living (NADED 249999). This value is the

number of diatoms valves of each taxa estimated to have originated from

living diatoms in the sample.

• Cell Density Factor. This is a duplication of the CDF from the Palmer-

Maloney and Diatom Slide Applied Factors Table – see above. This value

is derived for the sample as a whole and consequentially is the same for

each taxa in the sample.

• Benthic Cell Density. This computed value reflects the quantity of cells for

each taxa estimated to occur per cm2 of benthic surface. For each taxa

identified from the diatom slide, the Benthic Cell Density is equal to that

taxa’s Proportioned Living Diatom Valves multiplied by the sample CDF.

Whereas, for each taxa identified in the Palmer-Maloney slide, the Benthic

Cell Density is equal to Number of cells counted in the Palmer-Maloney

slide multiplied by the sample CDF.

• Biovolume per Cell – Applied Constants. This value will be incorporated by

DEP as part of its data upload into the DEP Algae Database. Estimated

biovolumes per cell for each taxa type may be determined by two

procedures. One procedure is to make measurements on each algae cell

3-138

observed in the diatom slide or in the Palmer-Maloney chamber and

compute an average biovolume per cell for each taxa. The procedure for

calculation may follow that offered by the Academy of Natural Sciences,

which is available at http://diatom.ansp.org/nawqa/Biovol2001.aspx. The

benefit of this procedure is that the biovolumes used are more reflective of

the sample. However, the difficulty with the process of making

measurements of every cell observed, or even a subsampling of those

observed, is that a large amount of time may be expended to obtain such

measurements. A less time consumptive procedure is to use archived

taxa-level biovolume data from the Academy of Natural Sciences. This

archived data is available from

https://diatom.ansp.org/autecology/download.asp?file id=4 and provides

various biovolume statistics for each taxon. These biovolume statistics are

compiled from the data the Academy has accumulated over many years of

research and would provide a reliable representation of the sampled taxa

population. When relying upon this later procedure, DEP would typically

use the average biovolume data statistic for each taxon, although other

values such as minimum, and maximum biovolumes are also available.

The problems with this alternative procedure are that it is not sample

specific and the data offered on the Academy website was last updated in

2001. This Academy provided biovolume data is further complicated by

the fact that many taxa names have changed since 2001 and many new

taxa may have since be added to this now dated list. Because of possible

taxa name changes, synonyms may have to be researched to determine

the biovolume of a taxon from this dated Academy list. However, these

problems could be partially solved by simply referring to that biovolume

data already existing as part of DEP Algae Database from samples

collected and processed by The Academy in 2013, 2014, 2015, and 2016.

At the beginning of each project, DEP will stipulate to the PL which of

these two procedures are to be used. [Note: connections to both of the

mentioned web sources may also be found at The Academy Autecology

site found at https://diatom.ansp.org/autecology/#browse.]

• Benthic Cell Biovolume. This historically PL reported value is presently not

included in this table exchanged between DEP and the PL. This value will

be incorporated by DEP as part of its data upload into the DEP Algae

Database. This computed value is equal to the Benthic Cell Density

multiplied by Biovolume per Cell-Applied Constant.

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Table 4. Example of the Identification/Enumeration Results Table, limited to a single sample collected in 2013 (table

submitted by PL to DEP would contain results for all samples). The left-most column is not part of the actual Access or

Excel based Result Table but is provided here to demonstrate the analysis process used to generate the data.

PADEP Algae Sample

GIS Key ID

Taxonomic

NADED ID

Number

Algae Taxon Name with Author Reference

Natuaral Units

Counted in

Palmer-Maloney

Number

Cells

Counted in

Palmer-

Maloney

Number

Valves

Counted in

Diatom

Slide

Total

Number

Valves

Counted on

Datom

Slide

Diatom Valve

Relative

Abundance

Number

Living

Diatoms

Counted in

Palmer-

Maloney

Proportioned

Living Diatom

Valves

CDF - Cell

Density Factor

Benthic Cell

Density (cells per

cm2 of sampled

benthic surface

area)

20130424-0800-jbutt 1023 Achnanthidium pyrenaicum (Hustedt) Kobayashi 435 631 0.6894 291 200.61 8174.50 1639887.61

20130424-0800-jbutt 7043 Amphora pediculus (Kützing) Grunow 46 631 0.0729 291 21.21 8174.50 173413.40

20130424-0800-jbutt 16004 Cocconeis placentula Ehrenberg 2 631 0.0032 291 0.92 8174.50 7539.71

20130424-0800-jbutt 16005Cocconeis placentula var. euglypta (Ehrenberg)

Grunow1 631 0.0016 291 0.46 8174.50 3769.86

20130424-0800-jbutt 16011 Cocconeis pediculus Ehrenberg 12 631 0.0190 291 5.53 8174.50 45238.28

20130424-0800-jbutt 27008 Diatoma moniliformis Kützing 1 631 0.0016 291 0.46 8174.50 3769.86

20130424-0800-jbutt 27013 Diatoma vulgaris Bory 7 631 0.0111 291 3.23 8174.50 26389.00

20130424-0800-jbutt 34030 Fragilaria vaucheriae (Kützing) Petersen 4 631 0.0063 291 1.84 8174.50 15079.43

20130424-0800-jbutt 37990 Gomphonema sp. 1 ? 6 631 0.0095 291 2.77 8174.50 22619.14

20130424-0800-jbutt 44073 Melosira varians Agardh 9 631 0.0143 291 4.15 8174.50 33928.71

20130424-0800-jbutt 45001 Meridion circulare (Greville) Agardh 1 631 0.0016 291 0.46 8174.50 3769.86

20130424-0800-jbutt 46023 Navicula gregaria Donkin 5 631 0.0079 291 2.31 8174.50 18849.28

20130424-0800-jbutt 46104 Navicula tripunctata (Müller) Bory 1 631 0.0016 291 0.46 8174.50 3769.86

20130424-0800-jbutt 46154 Navicula rhynchocephala Kützing 2 631 0.0032 291 0.92 8174.50 7539.71

20130424-0800-jbutt 46646 Navicula caterva Hohn et Hellerman 2 631 0.0032 291 0.92 8174.50 7539.71

20130424-0800-jbutt 46661 Navicula capitatoradiata Germain 2 631 0.0032 291 0.92 8174.50 7539.71

20130424-0800-jbutt 46859 Navicula lanceolata (Agardh) Kützing 10 631 0.0158 291 4.61 8174.50 37698.57

20130424-0800-jbutt 46893 Navicula antonii Lange-Bertalot 4 631 0.0063 291 1.84 8174.50 15079.43

20130424-0800-jbutt 46896 Navicula rostellata Kützing 2 631 0.0032 291 0.92 8174.50 7539.71

20130424-0800-jbutt 48002 Nitzschia acicularis (Kützing) Smith 2 631 0.0032 291 0.92 8174.50 7539.71

20130424-0800-jbutt 48004 Nitzschia amphibia Grunow 1 631 0.0016 291 0.46 8174.50 3769.86

20130424-0800-jbutt 48008 Nitzschia dissipata (Kützing) Grunow 22 631 0.0349 291 10.15 8174.50 82936.84

20130424-0800-jbutt 48013 Nitzschia frustulum (Kützing) Grunow 21 631 0.0333 291 9.68 8174.50 79166.99

20130424-0800-jbutt 48025 Nitzschia palea (Kützing) Smith 2 631 0.0032 291 0.92 8174.50 7539.71

20130424-0800-jbutt 48123 Nitzschia pusilla Grunow 1 631 0.0016 291 0.46 8174.50 3769.86

20130424-0800-jbutt 55002 Reimeria sinuata (Gregory) Kociolek et Stoermer 11 631 0.0174 291 5.07 8174.50 41468.42

20130424-0800-jbutt 57002 Rhoicosphenia abbreviata (Agardh) Lange-Bertalot 9 631 0.0143 291 4.15 8174.50 33928.71

20130424-0800-jbutt 65048 Surirella minuta Brébisson 3 631 0.0048 291 1.38 8174.50 11309.57

20130424-0800-jbutt 110004 Encyonema minutum (Hilse) Mann 4 631 0.0063 291 1.84 8174.50 15079.43

20130424-0800-jbutt 155003Planothidium lanceolatum (Brébisson ex Kützing)

Lange-Bertalot2 631 0.0032 291 0.92 8174.50 7539.71

20130424-0800-jbutt 155017Planothidium frequentissimum (Lange-Bertalot)

Lange-Bertalot1 631 0.0016 291 0.46 8174.50 3769.86

20130424-0800-jbutt 249998 (undifferentiated) diatoms> (dead) 53 53 8174.50 433248.50

20130424-0800-jbutt 249999 (undifferentiated) (diatoms)>(living) 291 291 8174.50 2378779.50

20130424-0800-jbutt 309000 Cladophora glomerata (Linnaeus) Kützing 3 11 8174.50 89919.50

20130424-0800-jbutt 510001 Scenedesmus quadricauda (Turpin) Brébisson 1 4 8174.50 32698.00

20130424-0800-jbutt 533000 Spirogyra sp. 1 4 8174.50 32698.00

20130424-0800-jbutt 569000 Ulothrix sp. 5 52 8174.50 425074.00

20130424-0800-jbutt 893002 Pleurocapsa minor Hansgirg 5 95 8174.50 776577.50

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Charles, D. F., C. Knowles, and R. S. Davis. 2002. Protocols for the analysis of algal

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Sabater, H. van Dam, and J. Vizinet. 1998. Recommendations for the routine

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in the Columbia basin, southeastern Washington. Ecology 39(1):46–65.

Wolda, H. 1981. Similarity indices, sample size and diversity. Oecologia 50:296–302.

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3.11 BACTERIOLOGICAL DATA COLLECTION PROTOCOL

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Prepared by:

Megan Bradburn and Shawn Miller






2015

Edited by:

Josh Lookenbill






2017

Edited by:

Shawn Miller






2021

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INTRODUCTION

All waters of Pennsylvania are evaluated for Water Contact Sports (WC) recreational

use (RU) attainment according to the criterion for bacteria in §93.7 of 25 Pa. Code

Chapter 93. This protocol identifies appropriate sample collection procedures needed to

assess for WC RU for applicable waters.

SAMPLING DESIGN CONSIDERATIONS SPECIFIC TO BACTERIA MONITORING

Probability-based or targeted (judgmental) sampling designs can both be used for

bacteria monitoring; however, DEP prefers a targeted sampling design. Both sampling

design procedures are described in more detail in Chapter 2, Sampling Design and

Planning (Lookenbill 2020). Watersheds that have not been assessed and waters that

are heavily used for recreational activities by the public or those that support public

beaches are prioritized and specifically targeted for recreational use monitoring.

Additional sampling locations are chosen throughout a watershed based on changes in

landuse, location of major tributaries, and potential sources of impairment. Impairment

sources can include municipal point sources, combined sewer overflows (CSOs), and

agricultural sources relating to manure application, livestock grazing, and animal

feeding. In most cases multiple sampling locations will be needed to bracket potential

sources of impairment. Smaller headwater streams (Strahler 2nd order or smaller),

especially those with no potential sources of impairment are not prioritized. For larger

rivers, streams, or lakes multiple sampling locations across a transect and along the

shore (cross-section surveys) are recommended to accurately delineate an

assessment. Cross-section surveys and multiple sampling locations across a transect

are highly recommended to be established on rivers and streams greater than 1000 mi2.

Cross-section surveys performed using a clean and calibrated field meter that collects

water temperature, specific conductance, dissolved oxygen, pH, and – preferably –

turbidity are required to determine if major water quality influences exist at the sampling

location according to Chapter 4.1, In-Situ Field Meter and Transect Data Collection

Protocol (Hoger 2020). The number and distribution of docks, boating areas,

residences, cabins, and access areas should be considered in locating sample sites.

Lakes and reservoirs with beaches or swimming areas must include sampling locations

from the left perimeter, center, and right perimeter of the swimming area, unless the

swimming area is less than 100 feet. If less than 100 feet, sampling locations from the

left perimeter and right perimeter are sufficient.

During the swimming season, the DOH and DCNR collect weekly samples for

Escherichia coli (E. coli) at public beaches for monitoring purposes. See 28 Pa. Code

§18.30. When exceedances of criteria occur at public beaches, closure notices are

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issued by DOH. In cooperation with DEP, DOH, and DCNR compiles a list of closures

that DEP will utilize to identify possible recreational use impairment and focus future

bacteriological data collection and WC RU assessment efforts.

SAFETY PROCEDURES

Appropriate WC RU assessment begins with data collection that follows proper

guidelines and safety procedures. Safety is of utmost concern when working in and on

any waterbody. Individuals that lack experience and education related to navigating

these waterbodies put themselves and others at risk of injury or loss of life. On large

waterbodies some form of watercraft can be required for this data collection protocol.

Whether a kayak, canoe, or powered boat is used, DEP recommends – and in most

cases PFBC requires by law – that individuals using watercrafts take boating courses

and obtain a boating safety certificate. However, whether laws apply to certain

individuals or watercraft or not, DEP strongly recommends that all individuals regardless

of age, experience, or navigation procedure obtain and keep a boating safety

certification on them during surveys that require watercraft. More information on boating

courses and safety certifications are available here:

http://www.fishandboat.com/Boat/BoatingCourses/Pages/default.aspx

Surface waters present difficulties for both wadeable and non-wadeable water

navigation. Certain locations, especially on large rivers, may be fully or partially

wadeable during certain times of the targeted recreation season, but there are also

highly variable currents and drop-offs that are dangerous to wading collectors.

Inherent in these recommendations is the understanding that collectors must be well

informed of the unique characteristics of each surface water they intend to sample and

be aware of current flow and weather conditions. Equipment considerations may be

different between similar sized surface waters, and certainly different when visiting

multiple surface waters of different size and type. For instance, boat selection and

safety equipment lists would be different depending on whether the collector is visiting a

small 3rd order stream or a large 7th order river. These differing characteristics require

thorough planning before going into the field so resources are not wasted due to a lack

of preparation. It is also imperative that collectors have access to and continually check

surface water flow conditions. There will be situations when weather in one part of

Pennsylvania may be optimal for sampling a site, but weather in another part of

Pennsylvania – two or three days before the sampling period – created conditions that

reduces the ability to collect. Analysis of flow conditions at all available points upstream

of the intended sampling locations are highly recommended. Commonly used discharge

data are available here: https://waterdata.usgs.gov/pa/nwis/current/?type=flow

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DATA COLLECTION

Bacteriological sampling for determining WC RU attainment is conducted during the

swimming season (May 1st through September 30th). This is the timeframe the public is

likely to be engaging in primary WC activities. WC recreational activities are defined

activities where immersion and ingestion are likely and there is a high degree of bodily

contact with the water, such as swimming, bathing, surfing, water skiing, tubing, skin

diving, snorkeling, water play by children, or similar water-contact activities (USEPA

2012). As a general rule, bacteriological sampling should not take place during those

times when physical conditions, such as stream discharge, render primary contact

recreation hazardous to the public. Samples collected for assessment purposes should

provide the ability to identify an impaired RU due to chronic long-term impacts and not

acute or transitory situations. In the past, there was guidance that sampling should not

take place immediately following 0.25 inch of rain or more to avoid collecting samples

during elevated discharge when bacteria levels are elevated, but not necessarily

characteristic of a chronic condition. This guidance is usually applicable to smaller

waterbodies where discharge responds quickly to precipitation events but becomes

more difficult to apply to larger waterbodies and to some lakes where increased

discharge and water quality changes can occur days following 0.25 inch of rain.

The WC RU bacteria criterion at 25 Pa. Code §93.7:

“(Escherichia coli colony forming unit per 100 milliliters (CFU per 100 ml)) During

the swimming season (May 1 through September 30), the maximum E. coli level

shall be a geometric mean of 126 CFU per 100 ml. The geometric mean for the

samples collected in the waterbody should not be greater than 126 CFU per 100

ml in any 30-day interval. There should not be greater than a 10% excursion

frequency of 410 CFU per 100 ml for the samples collected in the same 30-day

duration interval. (Fecal coliforms/100 ml) For the remainder of the year, the

maximum fecal coliform level shall be a geometric mean of 2,000 CFU per 100

ml based on a minimum of five consecutive samples collected on different days

during a 30-day period.”

The criterion specifically defines sample duration during the bathing season and

magnitude, frequency, and duration during the non-bathing season for determining

impairment for WC RU. The DEP Bacteriological Assessment Method for Water Contact

Sports, defines magnitude, frequency, and duration during the bathing season and

remainder of the year as at least 5 samples, each separated by at least 24 hours,

spanning a maximum of 30 days and a minimum of 14 days (Miller and Whiteash 2021).

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The frequency and duration of 5 samples over 30 days were included in USEPA’s

Recreational Water Quality Criteria Recommendations (2012):

“EPA believes that a shorter duration (i.e., 30 days), used in a static or rolling

manner, coupled with limited excursions above the [statistical threshold value]

STV, allows for the detection of transient fluctuations in water quality in a timely

manner. In the development of their monitoring program, EPA recommends that

states consider the number of samples evaluated in order to minimize the

possibility of incorrect use attainment decisions…”

Collecting data for water assessments in general will include samples collected

frequently enough and over a sufficient period of time to account for sampling error or

variations driven by confounding variables (Biggs and Whiteash 2021). WC RU samples

will include at least 5 bacteriological samples collected within a 30-day period.

Collecting samples throughout the entire May 1st through September 30th for surface

water assessment is not sustainable. However, it is recommended that additional

samples are collected to better understand the variability in the data and target

conditions that are likely to characterize impairment. This results in sampling error

decreases and a more confident assessment (Biggs and Whiteash 2021). If information

is available that indicates more than 5 samples over a 30-day period are required to

confidently assess, then additional samples will be collected.

DEP collects and analyzes DNA for Bacteroides dorei to identify the host animal of the

bacteria present in surface waters. B. dorei is a common bacterium found in the

environment and has a known genetic marker for the HF183 human genetic marker.

The process to analyze the DNA is known as quantitative Polymerase Chain Reaction

(qPCR) which results in genes being copied. qPCR is collected at sampling locations

that are suspected to have or have documented elevated levels of bacteria. DNA target

sequences are amplified in the presence of primers (short genetic sequences) that are

specific to various hosts (humans, cows, pigs, horses, birds, etc.). Results from qPCR

analyses are reported in units that are calculated based on the target DNA sequences

from test samples relative to those in calibrator samples that contain a known quantity

of target organisms (Haugland et al. 2005, Wade et al. 2010), which can aide in

determining the source of bacterial contamination.

In addition to collecting discrete bacteriological samples, continuous instream

monitoring (CIM) data can also be used to make assessment decisions. Water quality

data sondes record instream parameters at defined intervals that are sufficiently short to

be considered continuous. Bacteriological parameters are not capable of being directly

measured on a continuous basis, but models have been developed by comparing

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discrete bacteriological samples to recorded CIM data (Hoger 2018, Rasmussen et al.

2008, Stone and Graham 2014). Models may be used to collect water contact

recreational use data for assessment purposes, provide that CIM data collection

protocols are followed (Hoger et al. 2017).

COLLECTION REQUIREMENTS

Collector Identification Number

Collectors must have an assigned four-digit collector identification number (e.g., 0925).

This number along with a sequential three-digit sample number (e.g., 0925-001), and

date/time of sample are used to identify individual samples. Supervisory staff can

request collector identification numbers for their field staff with the “Collector ID Request

Form” found at the eLibrary website (ID Request Form).

DEP Laboratory Submission Sheet

Collectors must submit samples to the DEP Bureau of Labs (BOL) using the “Sample

Submission Sheet” (Submission Sheet). Field staff are required to document collector

identification number, reason code, cost center, program code, sequence number, date

collected, time collected (in military time format), fixative(s), Standard Analysis Code

(SAC), legal seal numbers for each sample collected (if required), the number of bottles

submitted per test suite, collector name, date, phone number, and any additional

comments that lab analysts will use to properly handle samples.

As described previously, collector identification numbers are unique to each field staff

collecting samples. Reason codes, cost centers, and program codes are program

specific and should be obtained from the program responsible for coordinating sampling

efforts. Sample sequence numbers are three-digit sequential numbers (001-999) unique

to a sample collected on a given day generated by field staff collecting samples. Date

and time collected should be accurately documented, especially if field parameters with

specific diurnal fluctuations (temperature, dissolved oxygen) will accompany analytical

results. To avoid complications with daylight savings time, collection time should be

recorded using eastern standard time or UTC-5 throughout the year.

A SAC is a unique code that details analytical tests to be applied to a specific sample.

Each DEP program uses specific SACs for specific projects or purposes. For example,

SAC B021 is used by “Water Management” when submitting bacteriological samples for

bot Fecal and E.coli with a 24-hour hold time. Unique SACs are created for specific

purposes. Table 1 lists other bacteriological test descriptions and SACs.

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Table 1. Bacteriological Test Descriptions and SACs

Organism Test Description SAC

Fecal coliforms

Membrane Filtration (8-hr. Hold

Time) B002

Fecal coliforms

Membrane Filtration (24-hr hold

time) B032

Fecal coliforms + Strep Membrane Filtration B003

Fecal + E.coli


time) B021

E. coli


time) B022

Enterococci Membrane Filtration B015

Fecal coliforms +

Total/Human/Avian/Bovine/Swine Bacteroides Membrane Filtration + qPCR B035

E. coli +

Total/Human/Avian/Bovine/Swine/Horse/Dog/Deer

Bacteroides Membrane Filtration + qPCR B037

Fecal + E. coli +

Total/Human/Avian/Bovine/Swine Bacteroides Membrane Filtration + qPCR B036

Total/Human/Avian/Bovine/Swine Bacteroides qPCR B038

Legal seals and associated legal seal numbers are required under circumstances where

it is imperative to document the integrity of samples from sample collection to sample

analysis. Legal seals are not always required and should be used per a program’s

specific requirements. Legal seal numbers must be singly listed (include letter and

number) for each sample. Legal seals can be obtained from BOL. Refer to BOL for

more information concerning legal seals.

Collector Name, Date, Phone Number, and # of Bottles submitted were added to the

laboratory submission sheet to meet NELAP chain-of-custody requirements. Each

submitted form is also required to have printed the collector’s name, the date, collector’s

signature (Relinquished by:), and collector’s phone number. BOL recommends

contacting the appropriate BOL staff before submitting samples.

Sampling Supplies and Equipment

• BOL Sample Submission Sheet (Submission Sheet)

• DEP Bacteria Monitoring Field and Site Data Sheets (Appendix A.10)

• List of sites, maps, and/or GPS

• 125-mL screw-capped, polypropylene or polystyrene bottles with sodium

thiosulfate

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• 1-L screw-capped, polypropylene bottles with sodium thiosulfate (qPCR)

• Sample blanks containing sterile (autoclaved) water obtained from BOL

• Disposable, powderless gloves (such as nitrile)

• Coolers and ice

• Courier shipping labels

• Van Dorn or Kemmerer (Lakes)

• Field meter (required for rivers and streams greater than 1000 mi2)

• Sampling pole and zip-ties (optional)

• Boots and waders (optional)

COLLECTION PROCEDURES

Collectors must ensure that the samples collected will be representative of the aquatic

system of interest. Interference of the sampling process must be minimized; collectors

must be alert to conditions that could compromise the integrity of a water sample. The

most common causes of sample interference during collection include poor sample-

handling and preservation techniques, input from atmospheric sources, and

contaminated equipment or reagents. Each sampling location needs to be selected and

sampled in a manner that minimizes bias caused by the collection process and that best

represents the intended environmental conditions at the time of sampling.

Before handling sample bottles, the collector should ensure his or her hands are clean

and not contaminated from sources such as food, coins, fuels, mud, insect repellent,

sunscreen, sweat, nicotine, etc. Alternatively, collectors should wear disposable,

powderless gloves (such as nitrile).

Labeling Bottles

While the minimum required information for samples submitted to BOL is the collector

number and sequential sample number, collectors may add additional information such

as date and time collected, general test(s) description. This will help prevent confusing

what bottles are for which tests and to ensure the sample is properly preserved.

Labeling should be done so that at least 1” of space is left at the top of the bottle to

allow BOL to apply lab labels.

Permanent marker will rub off bottles during collection and transport. So, clear packing

tape is wrapped around the bottle to protect the hand-written labels. Using ball-point

and other non-permanent ink pens must be avoided. BOL discourages the use of

masking tape. Collectors must complete the field and site data sheets (Appendix A.10)

for samples collected and sites visited.

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Flowing Waterbodies

For flowing waterbodies, bacteriological samples will be collected by direct surface

water sampling. Avoid sampling in eddies, pools, side-channels, or in tributary mixing

zones unless necessary due to site-specific or other environmental considerations. The

collector should face upstream, taking care to not alter flow patterns or disturb substrate

sediments upstream of where they will collect the sample. Collection bottles should be

inserted into the water column vertically, facing down to avoid inadvertently collecting

surface debris/films. For slow moving streams with easily disturbed sediment, the

collector should sample from the stream bank, boat, or bridge using a sampling

extension pole. If sampling from a boat, the collector should sample near the bow as the

boat moves upstream or faces upwind. In smaller waterbodies that do not require

multiple locations across a transect, samples are collected at mid-depth in the

approximate thalweg (the line defining the points along the length of a stream bed with

the greatest depth). The collector should remove the stopper/lid from the sampling

container just before sampling, taking care not to contaminate the cap, neck, or the

inside of the bottle with his or her fingers, wind-blown particles, precipitation, clothes,

body, or overhanging structures. DEP uses 125-mL or 1-L (qPCR) screw-capped bottles

with sodium thiosulfate added to neutralize the effects of residual chlorine. DO NOT

RINSE THE BOTTLE BEFORE SAMPLING. Fill the bottle to the shoulder leaving

headspace. Once the sample is collected and capped, the collector should rinse any

large amount of dirt or debris from the outside of the container.

Lakes and Reservoirs

Bacteriological samples will be collected using disposable, powderless gloves (such as

nitrile (elbow length preferred) from lakes. Samples are collected at a depth of one

meter when possible and at least a 0.3-meter depth at a minimum. Samples collected at

a depth of one meter can be collected using a sampling pole or dispensed from depth

specific water sampling equipment (e.g., Van Dorn, Kemmerer). Samples from a boat

are collected with motor off. The collector should remove the stopper/lid from the

sampling container just before sampling, taking care not to contaminate the cap, neck,

or the inside of the bottle with his or her fingers, wind-blown particles, precipitation,

clothes, body, or overhanging structures. DEP uses 125-mL or 1-L (qPCR) screw-

capped bottles with sodium thiosulfate added to neutralize the effects of residual

chlorine. DO NOT RINSE THE BOTTLE BEFORE SAMPLING. Fill the bottle to the

shoulder leaving headspace. Once the sample is collected and capped, the collector

should rinse any large amount of dirt or debris from the outside of the container.

Preservation

The collector should vertically insert bottles into a cooler, right-side up. The samples

should be cooled with cubed or crushed ice. A sufficient amount of ice should be added

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to the cooler to ensure samples remain at ≤6°C during overnight shipping. BOL

personnel will note whether samples were shipped properly. Improperly shipped

samples may be subject to a data release request. The “Sample Submission Sheet”

(Submission Sheet) should be filled out, inserted into a Ziploc® bag, and attached to the

inside of the cooler lid. Courier shipping labels should be printed out during ordering so

they can be attached to the top of the cooler lid during sample drop-off. Shipping labels

are secured to the cooler lip with two pieces of packing tape on the left and right side;

taping all sides of the label makes removal difficult for lab technicians.

Quality Assurance

For quality assurance purposes, sample blanks containing sterile (autoclaved) water

obtained from BOL or the laboratory analyzing samples should be submitted for every

20 samples for each sampling trip/day to determine whether contamination is occurring

in any part of the sample collection, handling, or preservation process. Sample blanks

are “collected” by transferring sterile water from container received from BOL to a

labeled 125-mL. A duplicate grab sample should also be collected every 20 samples or

for each sampling trip/day to gauge testing variability and potential sources of

contamination due to collection procedures. Duplicate samples are collected

simultaneously with the associated environmental sample, using identical sampling and

preservation procedures. Both sample blanks and sample duplicates are assigned

unique, sequential sample numbers. Label duplicate and blank samples in the same

manner as environmental samples. DO NOT IDENTIFY DUPLICATE OR BLANK ON

THE SAMPLE BOTTLE. Duplicates and blanks must be documented appropriately in

SIS under the ‘Comments/Quality Assurance’ tab and linked to a monitoring point.

Field Meters

Field meters provide the ability to collect in-situ data that is not available through grab

samples submitted to BOL. Standard field parameters include dissolved oxygen,

temperature, specific conductance, pH, and turbidity. Collection of field meter data is

recommended for bacteriological sampling and required for rivers and streams greater

than 1000 mi2. See Chapter 4.1, In-Situ Field Meter and Transect Data Collection

Protocol (Hoger 2020) for more information on the use and proper calibration of field

meters.

Sample Holding Times

Water samples need to be shipped or delivered to BOL as soon as possible. The

collector should understand that certain laboratory analyses have “holding times” during

which tests must be conducted for result validity. Bacteria samples collected and

submitted for compliance purposes have a hold time of 8 hours. A hold time of 24 hours

may be applied to samples collected and submitted for WC RU assessment purposes.

3-154

Shipping Samples

All DEP district and regional offices are designated pick-up locations for water samples;

the samples must be dropped off for pick up by 1600 hours. Other locations exist, such

as at some Pennsylvania Department of Transportation facilities and some private

businesses, but these drop-off locations may require call-ahead notice to the current

courier, as they may not be visited daily. Further, the drop-off locations may require a

drop-off specific key to open the drop-off entrance lock. Drop-off locations are available

on the BOL website.

SAMPLE INFORMATION SYSTEM DOCUMENTATION

General guidelines for data entry are provided in Chapter 4.6, Sample Information

System (SIS) Data Entry Protocol (Shull 2017). Step-by-step processes of SIS data

entry are also provided in Appendix B.3. However, the following includes

recommendations specific to managing projects, monitoring points, and samples using

bacteriological data.

Projects

Create a project specific to each recreational use monitoring and assessment effort.

This will include delineation of subbasin targeted for a specific project. ‘Project ID’

should be formatted to include the letters ‘BAC’, four alpha characters, followed by two-

digit year. ‘Description’ should include four-digit year, “Bacteria Monitoring Project”,

followed by the targeted subbasin or watershed. Include in the ‘Comments’ a general

description of the purpose and scope of the project along with any pertinent specific

information (Figure 1)

3-155

Figure 1. SIS Project entry and edit screen with template information.

Monitoring Points

Create a monitoring point for each sampling location. The ‘Name’ of each monitoring

point will begin with the same four alpha characters followed by sequential numbers.

Within the location tab populate ‘State’, ‘County’, ‘Latitude’, ‘Longitude’, ‘Datum’

(NAD83), and ‘Location’. Include a general location description in ‘Location’ (Figure 2).

Within the Function/Administration tab ‘Medium Type’ should be “Water” and for surface

water samples ‘Sample Medium’ will be “Surface Water” (Figure 3). Within the Aliases

tab ‘Alias ID’ will begin with the same four alpha characters followed by sequential

numbers as ‘Name’. Description will include the watershed, year, and targeted site

number (Figure 4). Within the NHD tab click ‘Launch NHD Locator, snap the monitoring

point to a NHD reach, and ‘Get NHD. Lastly, link the Monitoring Point to the Project by

clicking ‘Projects’, populating the ‘Project ID’, ‘Alias ID’, and ‘Effective Date’ (Figure 5).

3-156

Figure 2. SIS Monitoring Point, Location tab entry and edit screen with template

information.

Figure 3. SIS Monitoring Point, Function/Administration tab entry and edit screen with

template information.

3-157

Figure 4. SIS Monitoring Point, Aliases tab entry and edit screen with template

information.

Figure 5. SIS Monitoring Point, Project link and edit screen with template information.

3-158

LITERATURE CITED

Biggs, H., and R. Whiteash. 2021 (editors). Discrete physicochemical assessment

method. Chapter 3.1, pages 2–13 in D. R. Shull, and R. Whiteash (editors).

Assessment methodology for streams and rivers. Pennsylvania Department of


Miller, S., and R. Whiteash. 2021 (editors). Bacteriological assessment method for

water contact sports. Chapter 2.5, pages 75–79 in D. R. Shull, and R. Whiteash



Haugland, R. A., S. C. Siefring, L. J. Wymer, K. P. Brenner, and A. Durfour. 2005.

Comparison of Enterococcus measurements in fresh water at two recreational

beaches by quantitative polymerase chain reaction and membrane filter culture

analysis. Water Research 39:559–568.





Hoger, M. S., D. R. Shull, and M. J. Lookenbill. 2017. Continuous physicochemical data




Pennsylvania.

Hoger, M. S. 2018. Continuous physiochemical assessment method. Chapter 3.2,

pages 14–34 in D. R. Shull, and R. Whiteash (editors). Assessment methodology

for streams and rivers. Pennsylvania Department of Environmental Protection,






Rasmussen, T. J., Lee, C. J., and Ziegler, A. C. 2008. Estimation of constituent

concentrations, loads, and yields in streams of Johnson County, northeast

Kansas, using continuous water-quality monitoring and regression models,

October 2002 through December 2006: U.S. Geological Survey Scientific

Investigations Report 2008–5014. U.S. Geological Survey, Kansas Water

Science Center, Lawrence, Kansas. (Available online at:

https://pubs.er.usgs.gov/publication/sir20085014.)

3-159





Stone, M. L., and J. L. Graham. 2014. Model documentation for relations between

continuous real-time and discrete water-quality constituents in Indian Creek,

Johnson County, Kansas, June 2004 through May 2013: U.S. Geological Survey

Open-File Report 2014–1170. U.S. Geological Survey, Reston, VA. (Available

online at: https://pubs.usgs.gov/of/2013/1014/of13-1014.pdf.)

USEPA. 2012. Recreational water quality criteria. EPA 820-F12-058. Environmental

Protection Agency, Office of Environmental Information, Washington, DC.

Wade, T. J., E. Sams, K. P. Brenner, R. Haugland, E. Chern, M. Beach, L. Wymer, C.

C. Rankin, D. Love, Q. Li, R. Noble, and A. P. Dufour. 2010. Rapidly measured

indicators of recreational water quality and swimming-associated illness at

marine beaches: A prospective cohort study. Environmental Health 9:66.

4-2

4.1 IN-SITU FIELD METER AND TRANSECT DATA COLLECTION PROTOCOL

4-3

Prepared by:

Mark Hoger






2017

Edited by:

Mark Hoger






2020

4-4

INTRODUCTION

Water quality instruments provide the opportunity to quickly gather current water quality

data but proper care and calibration of the equipment, and QA/QC documentation are

required for these data to be accurate and defensible. Many water quality meters have

interchangeable probes to record a wide variety of parameters. The most common

parameters are temperature, conductivity, pH, dissolved oxygen, turbidity, and depth. A

full discussion of probe technology and considerations for their selection and use is

provided in Chapter 4.3, Continuous Physiochemical Data Collection Protocol (Hoger et

al. 2017).

CALIBRATION

Proper calibration should follow manufacturers’ guidelines. In addition, DEP strongly

recommends several procedures to further insure reliability and defensibility of data. At

a minimum, meters should have calibration checked before each day of use. In most

circumstances, it is recommended that the sensors are calibrated at this time. All

calibration activity, including checks, need to be logged and records maintained for

quality assurance. A calibration form is provided in Appendix B.1.

Calibration for all parameters should bracket any expected values. A summary of

calibration recommendations is provided in Table 1. Before calibration, the sensors

should be rinsed three times with the calibration solution. While other parameters can

be calibrated before going out into the field, dissolved oxygen (DO) sensors should be

calibrated onsite to avoid any influence of changes in barometric pressure from changes

in altitude, weather, or pressurization in buildings. It is also recommended that DO

calibration be checked periodically throughout the day. Accurate turbidity calibration can

be more challenging than other parameters as the sensors are very sensitive to

standard contamination and light infiltration. Clean, dedicated sensor guards and

calibration cups should be used. In addition, when calibrating in zero standard, or any

other very low standard, it can be necessary to have the guard in place and use an

opaque container to submerge the sensor in the standard. This procedure more closely

replicates the deployed environment than calibrating in just the calibration cup, and

therefore, can provide more consistent and accurate calibration.

4-5

Table 1. Summary of calibration recommendations

Parameter Calibration Notes

Temperature Check against NIST certified

equipment

Specific

Conductance

One-point calibration with

checks

• Calibrate at a high value

(1000 µS/cm)

• Check at a low value (100

µS/cm)

• Rinse with Deionized (DI)

water and check in air

(should be 0 µS/cm)

If values above 1000 µS/cm are

expected, also check in a higher

standard.

pH Three-point calibration (4,7,10) Two-point calibration acceptable

if conditions are known

Dissolved

Oxygen

100% water-saturated air:

• Water in bottom of

calibration cup (do not

submerge sensor)

• Calibration cup on loosely

to prevent pressure build-

up

• Run a wipe cycle, if

possible, to remove water

droplets on sensor

• Calibration onsite to avoid

influence of barometric

pressure differences.

• Check throughout the day.

• Allow plenty of time in cold

weather.

Turbidity Three-point calibration (0, ~100,

~1000)

• If only two-point calibration

available, calibrate at 0 and

high value, check in low

value.

• Pour standards slowly to

avoid bubbles.

• Use opaque calibration cup

or shield from direct light.

4-6

CLEANING AND MAINTENANCE

Water quality instruments dedicated to periodic field meter use generally do not require

an intensive cleaning regiment. Sensors should be visually inspected during calibration

and any deposits or biological growth removed following manufacturers’

recommendations. An occasional cleaning with mild soap and water is recommended.

With proper maintenance and care, many sensors have a life-span of several years.

One exception is pH probes that have a limited supply of reference solution. Some

manufacturers provide a refillable reservoir of reference solution to prolong the life of

their pH probes. Those that do not have a refillable reservoir often need to be replaced

yearly. The pH millivolt readings should be recorded during calibration to track the life of

the probe. Additional information on cleaning and maintenance is provided by Hoger et

al. (2017) and the equipment’s manufacturer.

STORAGE

Most probes require being stored in a moist environment. A small amount of fluid in the

bottom of the calibration cup is sufficient. Submerging pH probes or other sensors with

a reference solution will deplete that solution and reduce the life of the sensor.

RECORDING

Care should be taken to allow meters to stabilize before any readings are recorded.

Temperature and conductivity should often stabilize within seconds. Depending on the

environment and life of the probe, other sensors like pH or DO may take a couple

minutes to fully stabilize. If necessary, leave the meter in situ while other tasks are

completed, so all parameters have plenty of time to stabilize.

Division of Water Quality stores field meter readings in the DEP’s Samples Information

System (SIS). Discrete readings at continuous instream monitoring (CIM) stations are

additionally stored with the CIM data as field visits in Aquarius.

CROSS-SECTION SURVEYS

Waterbodies are dynamic systems changing throughout time and space. Regular cross-

section surveys help determine how homogenous a system is, allow for better

understanding of the system as a whole, and lead to better study design and

management decisions. The confluence of a tributary, groundwater infiltration, point-

source discharges, and differences in flow, depth, and growth of photosynthetic

organisms can all result in inconsistent water quality. In general, wide, flat, slow-moving

4-7

streams are more heterogeneous. As stream channels narrow and slope increases,

better mixing often occurs. DEP frequently utilizes cross-section surveys to determine

proper deployment of continuous monitoring equipment and direct the collection of

biological and chemical samples.

As with any water quality meter work, DEP requires daily calibration (or calibration

checks) of the meter before performing cross-section surveys. In addition, post-survey

calibration checks are completed to confirm that any variation in the survey was not due

to calibration drift of the instrument.

Cross-section surveys should be conducted multiple times to document differences (or

similarities) over time. DEP refers to these repeated survey points as a water quality

transect. Established transects are often repeated monthly, but more importantly should

target various flow levels and other events that may lead to variation in the survey.

The number of measurement points in a transect is influenced by the stream size and

heterogeneity. In the smallest streams (width < 15m), surveys usually consist of three

points: left-descending bank (LDB), mid-channel, and right-descending bank (RDB).

Streams 15m to 50m wide often have five points: LDB, ¼ across, mid-channel, ¾

across, and RDB. As streams become wider than 50m, surveys are established at 10-

20m intervals. However, in Pennsylvania’s largest rivers, widths can reach over one

mile across; and in these cases, intervals may be in excess of 100m. These large-river

transects frequently use bridge piers or other reference points to facilitate consistency of

subsequent surveys. In the absence of stable reference points, a range finder is used to

measure distance to the bank.

Evaluation of Transect Data

When evaluating water quality transect data, compare each point to a reference point in

the transect. The reference point can be a point of interest in the transect (e.g., CIM

location, grab sample location) or just a point from the transect that is likely to be well

representative of the waterbody (e.g., mid-point). The difference between each point

and the reference point is then evaluated for significance. For temperature and pH, the

difference in readings between the reference point and a transect point is considered

significant if the difference is greater than 0.5 units. For specific conductance, DO, and

turbidity, the difference is considered significant if it is greater than 10% of the reference

point reading. If transects are conducted when turbidity is low (less than 10 FNU), a

difference of one FNU is equivalent to a 10% difference.

It is important to evaluate transects over a range of conditions to best understand shifts

in mixing patterns under various conditions. Most streams will show significant

4-8

differences in water quality in at least small sections of a transect for short periods of

time (e.g., along the bank, during a rain event). Transect evaluations should

characterize the extent of these spatial and temporal fluctuations.

Display and Storage of Transect Data

Properly collected transects, under a variety of water quality conditions, are likely to

encompass a wide range of values. Because of this, charting the raw values together

often leads to difficulty in discerning meaningful differences. For example, it would be

difficult to see a 0.5°C difference in temperature when charting together a summer

transect with water temperatures around 30°C and a winter transect with water

temperatures around 3°C. Therefore, transect data are best displayed by charting the

comparisons of transect points and the reference point. The thresholds of significance

should be included in the chart to facilitate a quick evaluation of the data.

All transect survey data are recorded and archived along with meter calibration

documentation. A sample transect form is provided in Appendix B.2. Measurement data

from established water quality transects are entered and stored in the SIS database.

More information on SIS is found in Chapter 4.6, Sample Information System (SIS) Data

Entry Protocol (Shull 2017).

LITERATURE CITED





Pennsylvania.





4-9

4.2 DISCRETE WATER CHEMISTRY DATA COLLECTION PROTOCOL

4-10

Prepared by:

Dustin Shull






2013

Edited by:

Dustin Shull






2017

Disclaimer:

The mention of specific trade names or commercial products does not constitute

endorsement or recommendation for use.

4-11

INTRODUCTION

As part of its water quality monitoring programs, DEP collects water chemistry data to

assess the quality of Pennsylvania’s surface water resources. This information is used

to detect or confirm pollution sources and causes, for routine water quality monitoring,

and to establish abiotic-biotic relationships. This document provides guidelines for

standardized water chemistry collection from surface waters. The procedures described

here are adapted from scientific, peer-reviewed procedures, and were developed by

technical experts. This protocol does not attempt to describe the entire spectrum of

water quality data-collection techniques (such as continuous instream monitors), but

does describe the surface water sampling procedures appropriate for typical DEP

investigations and may require modification as situations dictate.

Variations to this protocol will be dependent upon site conditions, equipment limitations,

or limitations imposed by the procedure. Investigators should document modifications

and report the final procedures employed. Investigators should be aware of, and work to

mitigate, the potential for sample contamination at all phases of the sample collection

process by observing proper sample collection, handling, and preservation procedures

described here. The most common sources of error (also known as “interference”) are

cross-contamination, poor sampling locations, and incorrect preservation.



Field staff required to collect surface water samples must have an assigned four-digit

collector identification number (e.g., 0925). This number along with a sequential three-

digit sample number (e.g., 0925-001), and date/time of sample are used to identify

individual samples. Supervisory staff will request collector identification numbers for

their field staff with the “Collector ID Request Form” found at the eLibrary website (ID

Request Form).

DEP Laboratory Submission Sheet

Field staff will need to have at least Querying and Sample Entry security roles for the

program or business unit that they are collecting samples for. The program or business

unit will be consistent with the Program Code entered on the “DEP Laboratory

Submission Sheet” (Submission Sheet). Collectors must submit samples to the DEP

Bureau of Labs (BOL) using the laboratory submission sheet”. Field staff are required to

document collector identification number, reason code, cost center, program code,

sequence number, date collected, time collected (in military time format), fixative(s),

Standard Analysis Code (SAC), legal seal numbers for each sample collected (if

4-12

required), the number of bottles submitted per test suite, collector name, date, phone

number, and any additional comments that lab analysts will use to properly handle

samples.


collecting samples. Reason codes, cost centers, and program codes are program

specific and should be obtained from the program responsible for coordinating sampling

efforts. Sample sequence numbers are three-digit sequential numbers (001-999) unique

to a sample collected on a given day generated by field staff collecting samples. Date

and time collected should be accurately documented, especially if field parameters with

specific diurnal fluctuations (temperature, dissolved oxygen) will accompany analytical

results. To avoid complications with daylight savings time, collection time should be

recorded using eastern standard or UTC-5 time throughout the year. However, sample

holding times will need to be considered when there are time differentials.

A SAC is a unique code that details analytical tests to be applied to a specific sample.

Each DEP program uses specific SACs for specific projects or purposes. For example,

SAC 018 is used by collectors when submitting water chemistry samples for Special

Protection Surveys. The analytes/tests listed under SAC 018 are those specifically

identified by regulations that surface water must meet and therefore be assessed for, if

a special protection determination is warranted. Other programs have developed unique

SACs for their specific purposes, and the DEP BOL encourages programs to create

SACs tailored to a program’s specific needs. New SACs can be requested by filling out

the Request Form for Standard Analysis Code. An example of a commonly used SAC is

provided below (Table 1).

4-13

Table 1. Chemical parameters collected per SAC 612 during routine semi-wadeable

river surveys.

Test

Code Test Description

Reporting

Limit Units Reference

00095 Specific Conductance

at 25C 1 UMH/CM SM 2510B

00403 pH 0 pH UNITS SM 4500H-B

00410 Alkalinity, pH 4.5 0 MG/L SM 2320B

00530 Total Suspended

Solids 2 MG/L USGS-I-3765

00600A Total Nitrogen 0.064 MG/L SM 4500N-ORG

00602A Dissolved Nitrogen 0.064 MG/L SM 4500N-ORG

00608A Dissolved Ammonia 0.02 MG/L EPA 350.1

00610A Total Ammonia 0.02 MG/L EPA 350.1

00630A Total Nitrite and

Nitrate 0.04 MG/L EPA 353.2

00631A Dissolved Nitrite and

Nitrate 0.04 MG/L EPA 353.2

00665A Total Phosphorous 0.01 MG/L EPA 365.1

00666A Dissolved

Phosphorous 0.01 MG/L EPA 365.1

00671A Dissolved

Orthophosphate 0.01 MG/L EPA 365.1

00680 Total Organic Carbon 0.5 MG/L SM 5310C

00900 Total Hardness 0.11 MG/L SM 2340A+B and EPA 200.7

revision 4.4

00916A Total Calcium 30 UG/L EPA 200.7 Revision 4.4

00927A Total Magnesium 10 UG/L EPA 200.7 Revision 4.4

00929A Total Sodium 200 UG/L EPA 200.7 Revision 4.4

00937A Total Potassium 1 MG/L EPA 200.7 Revision 4.4

00940 Total Chloride 0.5 MG/L EPA 300.0

00945 Total Sulfate 1 MG/L EPA 300.0

01007A Total Barium 10 UG/L EPA 200.7 Revision 4.4

01022K Total Boron 200 UG/L EPA 200.7 Revision 4.4

01042H Total Copper 4 UG/L EPA 200.8 Revision 5.4

01045A Total Iron 20 UG/L EPA 200.7 Revision 4.4

01051H Total Lead 1 UG/L EPA 200.8 Revision 5.4

01055A Total Manganese 10 UG/L EPA 200.7 Revision 4.4

01067H Total Nickle 4 UG/L EPA 200.8 Revision 5.4

01082A Total Strontium 10 UG/L EPA 200.7 Revision 4.4

01092A Total Zinc 10 UG/L EPA 200.7 Revision 4.4

01105A Total Aluminum 200 UG/L EPA 200.7 Revision 4.4

01132A Total Lithium 25 UG/L EPA 200.7 Revision 4.4

01147H Total Selenium 7 UG/L EPA 200.8 Revision 5.4

70300 Total Dissolved Solids 2 MG/L USGS-I-1750

70507A Total Orthophosphate 0.01 MG/L EPA 365.1

82550 Osmotic Pressure 1 MOSM/KG DEP BOL

99020 Total Bromide 50 UG/L EPA 300.1

4-14



analysis. Legal seals are not always required, and should be used per a program’s

specific requirements. Legal seal numbers must be singly listed (include letter and

number) for each sample. Legal seals can be obtained from BOL. Refer to BOL for

more information concerning legal seals.

Collector Name, Date, Phone Number, and # of Bottles submitted were added to the

laboratory submission sheet to meet NELAP chain-of-custody requirements. Using the

area at the bottom of the form, each bottle submitted for the samples identified must be

accounted for by enumerating the number of bottles per category listed for inorganic

and organic analyses/tests. Each submitted form is also required to have printed the

collector’s name, the date, collector’s signature (Relinquished by:), and collector’s

phone number. There are also spaces to document a facility name, facility identification

number, and an alternate contact. These three pieces of information are not required.

The last piece of information to be documented is additional comments that lab analysts

will use to properly handle samples. This information is documented in the ‘Comment’

field at the bottom of the form. The most common use of this field is to add or delete

tests to or from a specified SAC. For example, SAC 018 does not include a test for

turbidity; however, a sample collector may want BOL to document turbidity for a sample

so they would indicate in the ‘Comment’ field to add the turbidity test to the sample. If

many samples will be submitted with consistent modifications of a SAC, the BOL prefers

a new SAC be created for those samples. Other important comments to consider

include identifying potentially toxic or otherwise dangerous samples, samples submitted

for individual tests, and other important information that lab analysts will need to know to

handle the samples correctly. The BOL recommends contacting the appropriate BOL

staff before submitting samples requesting organic tests, potentially dangerous

samples, or samples that need to be handled differently.


DEP programs can and do employ a multitude of program specific surface water

sampling techniques that will require standard supplies (e.g., sampling bottles,

preservatives, etc.), and specialized supplies (e.g., 0.10µm filters) or equipment (e.g.,

Van Dorn sampler). This section describes the equipment and supplies required to

collect the most commonly used surface water sampling techniques that may be

employed. Additional techniques are added as they become applicable and as standard

procedures are formalized.

4-15

Typically, a 500ml HDPE bottle is used for storing an unfiltered, non-chemically

preserved sample for inorganic constituent analyses and 125ml High-Density

Polyethylene (HDPE) bottles are used for storing filtered or unfiltered samples such as

metals and nutrients. Water samples collected for total or dissolved organic carbon or

volatile organic compound analyses are stored in two 40ml Volatile Organic Analysis

(VOA) amber glass vials. Additional analyses may require other specialized containers,

so it is important to check the BOL site when working with new analyses. A suggested

check list of recommended supplies and equipment is provided below:

4-16

SAMPLE CONTAINERS

500 ml sample bottles - inorganic, total metals, cyanides, phenolics, other

125 ml sample bottles - dissolved metals 1000 ml amber glass bottles - organics:

semi- volatiles, pesticides, PCBs 40 ml glass vials - organics: VOAs 125 ml bac-t’ bottles - bacteriological

analysis (fecal coliform & E.coli) other:

_______________________________ FIXATIVES

HNO3 H2SO4

pH test strips other: NaOH, HCl

FIELD METERS & RELATED SUPPLIES

dissolved oxygen meter DO probe solution zero % calibrating solution (if applicable)

pH meter buffers (pH 4, 7, 10) KCl probe storage solution

conductivity meter calibrating solution (if applicable)

thermometer meter field manuals (if applicable)

OTHER

0.45µm groundwater filters or appropriate filtration apparatus with filters

Alkalinity test kit Blank water (lab tested) Extra bottles for duplicates pipetter & pipettes buckets & rope (applicable length for

bridge sampling) rinse squirt bottle eyewash bottle

FLOW

flow meter rods (for anchoring tape bank-to-bank) tape measure wading rod

FORMS

laboratory water chem. sheets bac-t’ forms physical data field forms flow field form habitat assessment forms chlorine demand forms other:__________________

SHIPPING

courier shipping forms tape & dispenser shipping coolers ice

MISC.

maps GPS waders gloves markers, pens, & pencils calculator insect repellent screwdriver/tools batteries (D-cell, other:________) other:

_______________________________

4-17


Many surface water samples are manually collected instantaneous “grab” samples.

Water samples should be collected before conducting other types of in-stream field

work in the study reach, such as discharge measurements or benthic macroinvertebrate

collection to avoid disturbing the water column. Water samples are normally collected in

HDPE, VOA vials or amber glass bottles (for organic compound analysis). Samples

should be preserved immediately upon collection, if necessary, with the proper type and

amount of chemical fixative (usually an acid to an amount where the matrix pH is less

than 2.0 pH units). All sample bottles should be cooled and held at ≤6°C until receipt by

the laboratory. The sample bottles should be labeled in accordance with the procedure

described later in this document. The bottles used for a sample are determined by the

analyses required for that sample and the different types of preservation required for the

analyses of interest.

Some chemical analyses require laboratory technicians to calibrate specialized

laboratory equipment, prepare specialized reagents, or otherwise perform pre-analytical

preparation before samples can be analyzed. If a collector is going to submit several

samples involving specialized preparation (such as for bacteriological analyses, which

involve agar plating), they should contact the appropriate technician at the laboratory to

ensure enough time is allocated for the pre-test procedures. If the laboratory is not

notified to expect a large volume of these samples, holding times may be surpassed

and the sample may be voided.

Field personnel must ensure that the samples collected will be representative of the

aquatic system of interest. A grab sample temporally and spatially represents the part of

the surface water system being investigated. The data collection protocol and

constituents chosen for analysis is critically dependent on the purpose and scope of the

survey being conducted. Obtaining representative samples is of primary importance for

a relevant description of the aquatic environment. Interference of the sampling process

must be minimized; collectors must be alert to conditions that could compromise the

integrity of a water sample. The most common causes of sample interference during

collection include poor sample-handling and preservation techniques, input from

atmospheric sources, and contaminated equipment or reagents. Each sampling site

needs to be selected and sampled in a manner that minimizes bias caused by the

collection process and that best represents the intended environmental conditions at the

time of sampling.

Before handling sample bottles, the collector should ensure his or her hands are clean

and not contaminated from sources such as food, coins, fuels, mud, insect repellent,

4-18

sunscreen, sweat, nicotine, etc. Alternatively, collectors should wear disposable,

powderless gloves (such as nitrile).

Labeling Bottles

While the minimum required information is the collector number and sequential sample

number, collectors may add additional information such as date and time collected,

general test(s) description (total metals, TOC, etc.), “filtered”, and preservation type.

This will help prevent confusing what bottles are for which tests and to ensure the

sample is properly preserved. Labeling should be done so that at least 1” of space is left

at the top of the bottle to allow BOL to apply lab labels.

Permanent marker will rub off HDPE bottles during collection and transport. So, clear

packing tape is wrapped around the bottle to protect the hand-written labels. Using ball-

point and other non-permanent ink pens must be avoided. BOL discourages the use of

masking tape. Collectors should keep a log book of all samples they collect for later

reference. The sample log entry should annotate the unique collector identification and

sequence number, date and time, the waterbody name, sample location, SAC code,

and any additional analytical tests performed or excluded. Additional information on

labeling samples can be found on the BOL website.

Direct Surface Water Sampling of Wadeable, Flowing Waterbodies

The most common type of water sampling is conducted in wadeable, flowing

waterbodies, where water is collected directly into the sample bottle. This procedure is

not generally used in situations where contact with contaminants is a concern.

The collector should face upstream, taking care to not alter flow patterns or disturb

substrate sediments upstream of where they will collect the sample. Collection bottles

should be inserted into the water column vertically, facing down to avoid inadvertently

collecting surface debris/films. In most situations, samples are collected at mid-depth in

the approximate thalweg (the line defining the points along the length of a stream bed

with the greatest depth). The collector should remove the stopper/lid from the sampling

container just before sampling, taking care not to contaminate the cap, neck, or the

inside of the bottle with his or her fingers, wind-blown particles, precipitation, clothes,

body, or overhanging structures. All bottles are rinsed three times instream before filling

the bottle. Once the sample is collected and capped, the collector should rinse any dirt

or debris from the outside of the container.

Field Meters

Field meters provide the ability to collect in-situ data that is not available through grab

samples submitted to BOL. Standard field parameters include dissolved oxygen (DO),

temperature, specific conductance, pH, and turbidity. While BOL does report pH, it is

4-19

understood that the laboratory reported pH of a grab sample can and will migrate.

Therefore, the lab result may not reflect the actual instream pH. Collection of field meter

data is highly recommended and required depending on the specific field survey

protocol being used. Field meter data will be collected per Chapter 4.1, In-Situ Field

Meter and Transect Data Collection Protocol (Hoger 2020)

Collecting Unfiltered Samples

For unfiltered samples, the collection bottle is rinsed at least three times with the water

to be sampled. The collector removes the lid from the bottle and partially fills the bottle

under water. The bottle is then removed from the water, capped, shaken vigorously,

uncapped, and inverted. Rinsing waste is discarded downstream of the collector to

ensure no contamination reenters the sample bottle. Unfiltered, inorganic samples will

be filled to the neck of the bottle, allowing for “head space” as requested by the BOL.

Specialized bottles, such as VOAs and 1L amber glass jars, are not rinsed. VOAs used

to collect volatile samples, however, are filled to the top and capped so that no air

remains in the bottle. 1L Amber glass jars are submerged and filled only to the neck. 4L

cube containers (Cubitainers) are rinsed three times and filled. Regardless the bottle

type, it is imperative that the bottle is filled with water the same way every time to

maintain consistency. The proper amount of chemical preservative is then added (Table

2); the bottle is recapped and then inverted several times to mix the reagent with the

sample.

Collecting Dissolved (Filtered) Samples

Dissolved samples must be filtered and fixed immediately after sample collection using

a 0.45µm disposable (single-use, metals free) filter (e.g., AquaPrep 600 Groundwater

Filters, VWR: #28145-142 or equivalent). Filters should be rinsed with trace-metal free

deionized (“ultrapure”) water before sample collection begins. Rinsing removes trace

contaminants (if any) from the manufacturing process.

A 1000ml squeeze-type bottle or churn is typically used to collect the raw water to be

filtered. This container is rinsed three times in the same manner as for an unfiltered

sample. The filtering device (e.g., syringe) is also rinsed three times using the raw water

in the container before filtration begins. Raw water is then aliquoted from the container

into the filtering device and rinsed through the filter. Once all equipment is rinsed, raw

water may pass through the filter into the sample bottle. Again, the sample bottle must

be rinsed three times with filtered water before filling the sample bottle. As with the

unfiltered 125ml sample, water is filled to the neck of the bottle. It is imperative that the

collector accurately fills water to this line to ensure consistent dilution of the fixative

between samples. If the collector is not consistent in performing this quality assurance

step, his or her dissolved constituent concentrations may be greater than the equivalent

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total constituent concentration in the unfiltered sample. The reagent preservative is then

added and mixed.

Preservation

Without preservation, water sample constituents will continue chemical interactions or

undergo other physical processes, such as metals precipitation. Moreover, laboratory

pH measurements are usually higher than field measurements because of carbon

dioxide degassing from the matrix. Keeping the water samples ≤6°C minimizes these

processes. Most chemical preservatives function by decreasing the matrix’s pH below

2.0 (or above pH 12 for cyanide - fixed with NaOH), which limits constituent reaction.

For example, 125ml bottles require 2 ml of preservative to achieve a pH threshold <2.0

(Table 2). Most water samples measuring metals concentrations use 1:1 HNO3;

however, it is important to note that ferrous iron is preserved with 1:1 HCl, not HNO3.

The 10% H2SO4 solution is used for storing water samples measuring phosphorus and

nitrogen species such as ammonia. However, for VOA samples such as TOC, the 10%

H2SO4 solution should be obtained from the organics laboratory, because it is not the

same as standard 10% H2SO4 solution. If the recommended amount of preservative is

added then pH does not need to be checked after preservation. If a pH check is still

desired, then the testing device (pH test strip) is held below the sample bottle and a

small amount of the sample is poured on the device. The testing device should not

contact with the sample bottle.

Table 2. Recommended chemical preservative amount for commonly used sample

bottle sizes.

Bottle Size Preservative Amount

40ml VOA ≈ 1.0 ml

125ml HDPE 2.0 ml

500ml HDPE 8.0 ml

Reusing graduated pipettes to dispense preservatives is permitted, but the collector

should understand that doing so introduces potential for contamination at several points

in the preservation process. The pipette must be carefully guarded against contacting

any surface and should be replaced often. Additionally, the preservative contained in

the bulk storage container can become contaminated from constant opening and

insertion of the pipette. Therefore, it is recommended that preservatives and bulk

containers are refreshed regularly. Most preservatives are highly reactive acids that can

cause chemical burns to skin, reactive equipment, and clothing, so collectors should

exercise caution when handling, transporting and storing these chemicals. Refer to

material safety data sheets (MSDS) for proper safety precautionary procedures.

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Sample location considerations

In general, collectors should avoid sampling in eddies, pools, side-channels, or in

tributary mixing zones unless necessary due to site-specific or other environmental

considerations. Collectors must also be aware of potential point-source and non-point

sources of water quality influence.

For slow moving streams with easily disturbed sediment, the collector should sample

from the stream bank, boat, or bridge using a sampling extension pole. If sampling from

a boat, the collector should sample near the bow as the boat moves upstream or faces

upwind.

For point source surveys and characterizing other influences, representative water

samples are collected from the discharge pipe/influence, from upstream (control), and

downstream locations at a minimum. Sampling stations located upstream of the

discharge pipe should be in a non-impacted zone to serve as a control. If there are

multiple discharges, then sample stations should be placed to bracket individual

discharges to better characterize each source. When sampling downstream of the

discharge pipe, the investigator should avoid the immediate vicinity of the

discharge/influence point and select a sample point far enough downstream to allow for

mixing between the discharge and stream flow.

To decide where an acceptable downstream sampling point should be located, consider

the following. For pollutants controlled by acute concerns, enforcement of numeric

criteria is at the point of complete mix, or after 15 minutes of travel time, whichever

occurs first. For pollutants controlled by chronic concerns, enforcement of numeric

criteria is at the point of complete mix, or after 12 hours of travel time, whichever occurs

first. These are all projections at design conditions (Q7-10 or harmonic mean flow). The

actual point of complete-mix depends on the stream size, its width and depth, its flow on

that day, the velocity of the plume, the angle at which it enters the stream, and the

roughness of the stream bed.

Field specific conductance measurements may help determine the point of complete

mix. If the point of complete mix is unclear or too far downstream for representative

sampling, then multiple samples should be collected across the width of the waterbody.

For very large rivers it may be necessary to composite water samples collected along a

cross channel transect to accurately characterize water quality of the sampled stream

segment.

Before collecting data to assess the Potable Water Supply (PWS) use, collectors should

consider that while 25 Pa. Code § 93.4 and § 96.3(c) identify PWS as applying to all

surfaces waters, § 96.3(d) states,

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“(d) As an exception to subsection (c), the water quality criteria for total dissolved

solids, nitrite-nitrate nitrogen, phenolics, chloride, sulfate and fluoride established

for the protection of potable water supply [PWS] shall be met at least 99% of the

time at the point of all existing or planned surface potable water supply

withdrawals unless otherwise specified in this title”

For these parameters, the PWS use can be evaluated by collecting samples upstream

of the surface water withdrawal at a minimum of one location or from the facilities raw

water faucet that they use for their testing. However, multiple locations may be

necessary to identify potential sources of pollution. Analyses are performed for total

nitrites, total and dissolved metals, chloride, fluoride, sulfate, color, and dissolved solids

using SAC 166. Additional microbiological parameters can be added on a site-specific

basis.

Quality Assurance

For quality assurance purposes, sample blanks containing ultrapure water obtained

from BOL should be submitted for every 20 samples for each sampling trip/day to

determine whether contamination is occurring in any part of the sample collection,

handling, or preservation process. A duplicate grab sample should also be collected

every 20 samples or for each sampling trip/day to gauge testing variability and potential

sources of contamination due to collection procedures. Duplicate samples are collected

simultaneously with the associated environmental sample, using identical sampling and

preservation procedures. Both sample blanks and sample duplicates are assigned

unique, sequential sample numbers. The collector needs to carefully annotate which

sample is a duplicate or blank. Duplicates and blanks must be documented

appropriately in SIS under the ‘Comments/Quality Assurance’ tab.


Water samples need to be shipped or delivered to BOL as soon as possible. The

collector should understand that certain laboratory analyses have “holding times” during

which tests must be conducted for result validity. Nitrate concentrations, for example,

must be measured within 48 hours of sample collection. If a sample surpasses holding

time requirements, the results will not be reported unless a “Request to Analyze

Voidable Samples” form (see the BOL website) is submitted to the BOL. It is not

advisable to collect and ship samples on Fridays, as the laboratory does not operate on

weekends; samples shipped on Friday will not be received and logged until Monday

morning. Doing so will guarantee that holding times of 48 hours or less will not be met.

Collectors essentially need to plan their water sampling from Monday through Thursday,

dropping off samples collected those days by 1600 hours, and verify shipped samples

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will reach DEP BOL by early Friday morning at the latest. Samples collected for

Carbonaceous Biological Oxygen Demand (CBOD) and Biological Oxygen Demand

(BOD) analyses have a 48-hour holding time. Holding time begins at the time of

sample(s) collection. Initial DO is not performed on CBOD/BOD samples until

Wednesday due to the five-day incubation period; consequently, if samples must be

collected on Mondays, collect only after 1300 hours to ensure the test is within holding

time.

Shipping Samples

All DEP district and regional offices are designated pick-up locations for water samples;

the samples must be dropped off for pick up by 1600 hours. Other locations exist, such

as at some Pennsylvania Department of Transportation facilities and some private

businesses, but these drop-off locations may require call-ahead notice to the current

courier, as they may not be visited daily. Furthermore, some drop-off locations may

require a drop-off specific key to open the drop-off entrance lock, and access should be

arranged with the point of contact prior to sampling. Drop-off locations are available on

the BOL website.



to the cooler to ensure samples remain at ≤6°C during overnight shipping. Laboratory


samples may be subject to a data release request. Dry ice will freeze water samples

and should never be used for storage or shipping of water chemistry samples. The

“Sample Submission Sheet” should be filled out, inserted into a Ziploc® bag, and

attached to the inside of the cooler lid. Courier shipping labels should be printed out

during ordering so they can be attached to the top of the cooler lid during sample drop-

off. Shipping labels are secured to the cooler lip with two pieces of packing tape on the

left and right side; taping all sides of the label makes removal difficult for lab

technicians.

Other Sampling Considerations

For storm water surveys, a minimum of one sample is collected during low or dry

weather flow to determine background conditions and from 3 to 5 high flow (storm)

events in conjunction with stream flow measurements to characterize pollutant loadings.

For storm events, it is important to make collections during the first flush and/or while

the hydrograph is rising. Analyses should be performed for metals (Fe, Al, Cu, Pb, Zn,

Cd, Cr, Hg), oils and grease, pathogens, and for total and dissolved nutrients. Analysis

is not limited to the above, and parameters of special concern (e.g., fertilizers,

pesticides and other organic chemicals) may be added as necessary.

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If deemed necessary by the investigator, nutrient sampling will occur during the growing

season at least once a month from May through October. Sampling should occur during

both dry and wet weather to adequately characterize loadings. Wet weather samples

should be collected during the rising hydrograph. In addition, stream discharge will be

measured at least once. Water quality analyses should be conducted for total and

dissolved nutrients using SAC 047.

For abandoned mine discharges or acid mine discharges, samples should be collected

from the point(s) of discharge, if possible. In addition, flow from the discharge(s) should

be measured to determine loading rates for Total Maximum Daily Load (TMDL)

development. Flow and channel cross section are measured in the field per standard

USGS stream gauging techniques (USGS 2006). Analyses are performed for metals,

alkalinity, and acidity using SAC 909.

Atmospheric deposition sampling should occur in late winter/early spring during heavy

snowmelt and/or storm events to capture episodic acidification. Sampling should occur

during base flow and peak flow to characterize worst-case and document the difference

between base flow and worst-case conditions. This protocol includes a filtering

procedure for dissolved monomeric aluminum that differs from that prescribed for other

dissolved metals. Water for the dissolved aluminum analysis is filtered through a 0.1µm

filter rather than through the standard 0.45µm filter. A pump is recommended to aid this

filtration process. The result from this alternate dissolved aluminum analysis correlates

well with the occurrence of inorganic monomeric aluminum species, which causes lethal

responses in fish. Analyses are performed for metals, alkalinity and acidity using SAC

122, or as otherwise described in the corresponding assessment method (Shank 2021).

LITERATURE CITED





Shank, M. K. (editor). 2021. Atmospheric deposition source and cause determination

method. Chapter 6.2, pages 6–12 in D. R. Shull, and R. Whiteash (editors).

Assessment methodology for streams and rivers. Pennsylvania Department of


USGS. 2006. Collection methods at flowing-water and still-water sites. Book 9, chapter

A4, section 4.1.3 in National field manual for the collection of water quality data.

U.S. Geological Survey, Office of Water Quality, Reston, VA. (Available online at:

https://pubs.usgs.gov/twri/twri9a4/twri9a4 Chap4 v2.pdf.)

4-25

4.3 CONTINUOUS PHYSICOCHEMICAL DATA COLLECTION PROTOCOL

4-26

Prepared by:

Mark Hoger, Dustin Shull, and Josh Lookenbill






2017

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INTRODUCTION

DEP uses continuous instream monitors to assess the quality of Pennsylvania’s

streams. Data collected using continuous instream monitoring will be used in

conjunction with other data for evaluation of use attainment. Most instream monitoring

configurations include at least four parameters: water temperature, specific

conductance, pH, and dissolved oxygen. Monitors can also be configured to measure

additional properties, such as turbidity, water stage, and fluorescence. Sensor data are

valuable for a variety of purposes including but not limited to, characterizing baseline

physicochemical stream conditions, describing seasonal and diel fluctuations, and

documenting potential exceedances of water quality criteria. These data are also used

in conjunction with flow measurements and chemical analyses of grab samples to

estimate chemical loads. Sensors that are used to measure water quality field

parameters require careful field observation, cleaning, and calibration procedures. The

resulting data requires systematic actions for the computation and publication of final

records.

This protocol provides guidelines for site and monitor selection, sensor inspection and

calibration procedures, field maintenance, data evaluation and correction, and reporting

procedures. Many of these criteria and procedures were developed from and mirror the

USGS Guidelines and Standard Procedures for Continuous Water Quality Monitors:

Station Operation, Record Computation, and Data Reporting (Wagner et al. 2006).

Water quality parameters are dynamic, necessitating frequent and repeated

measurements to adequately characterize variations in quality. When the time interval

between repeated measurements is adequately small, the resulting water quality record

can be considered continuous. A device that measures water quality in this way is

called a continuous water quality monitor. These monitors have sensors and recording

systems to measure physicochemical water quality field parameters at discrete time

intervals and at discrete locations. Operation of a water quality monitor provides a

record of water quality that can be processed and reported. The water quality data

provides a record of changes in water quality that also serve as the basis for

computation of constituent loads at a given site. Data from the sensors are also used to

estimate other constituents if a significant correlation (typically through regression

analyses) can be established.

Water temperature, conductivity, dissolved oxygen (DO), pH, and turbidity are

commonly recorded using continuous monitors. Water temperature and conductivity are

true physical properties of waterbodies, whereas DO and pH are measured

concentrations. Turbidity is an expression of the optical properties of water. Additional

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sensors are available to measure other field parameters, such as oxidation-reduction

potential, water stage, ammonia, nitrate, chloride, and fluorescence. Some monitors

also include algorithms to report calculated parameters, such as specific conductance,

total dissolved solids, and percentage of DO saturation. Calculated parameters can be

useful; however, consideration for the potential error in the algorithms should be taken

into account. Sensor technology broadens the variety of measurable chemical

constituents and reduces the limits of detection. As a result, continual progress is being

made to improve applications and refine quality-control procedures.

The vast majority of continuous monitoring of water quality field parameters takes place

in Pennsylvania’s lotic surface waters which may vary significantly in size, clarity,

chemical composition, and biological production. Procedures for continuous monitoring

in pristine, headwater streams differ from those in large, impacted rivers. Continuous

monitoring in impacted environments can be challenging because of rapid biofouling

from microscopic and macroscopic organisms, corrosion of electronic components from

salts, and wide ranges in values of field parameters associated with changing weather.

This protocol provides basic guidelines and procedures for use by DEP personnel in

site/monitor selection, field maintenance and calibration of continuous water quality

monitors, record computation, review, and data reporting. This protocol details the

primary technique used by DEP for deploying and servicing continuous instream

monitors. This protocol highlights basic guidelines that may need to be modified to meet

local environmental conditions. Data management procedures provided in this protocol

are designed to offer basic guidelines and may need to be manipulated to suit other

purposes and different software. In-depth knowledge of equipment operation and

familiarity with the watershed are imperative in the data evaluation process. Examples

of the application of scientific judgment in the evaluation of data records are discussed

and are, by necessity, site specific.

MONITOR DEPLOYMENT

Major considerations in the design of a continuous instream monitoring station include

site selection, monitor selection, monitor configuration, and sensor selection. Sensor

and site selection are guided by the purpose of monitoring and the data objectives. The

main objective in the placement of the sensors is the selection of a stable, secure

location that is representative of the aquatic environment.

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Site Selection

The main factors to consider in selecting a water quality monitoring site are the purpose

of monitoring and the data quality objectives. All other factors used in the site selection

process must be balanced against these two key factors. Defining the purpose of

monitoring includes making decisions about the field parameters to be measured, the

period and duration of monitoring, and the frequency of data collection. More site-

specific considerations in monitor placement include site-design requirements, monitor-

installation type, physical constraints of the site, and servicing requirements.

Once the purposes of monitoring and data-quality objectives are defined, balancing the

numerous considerations for placement of a continuous water quality monitoring system

still can be difficult. Obtaining measurements representative of the waterbody usually is

an important data quality objective. The optimum site consideration for achieving this

objective is placing the sonde in a location that best represents the waterbody being

measured. Thus, an optimal site is one that permits sensors to be located at a point that

best represents the section of interest for the aquatic environment being monitored.

Cross-section variability and upstream influences are major factors for site selection.

Cross-section surveys of field parameters must be made to determine the most

representative location for monitor placement. A site must not be selected without first

determining that the data-quality objective for cross-section variability will be met.

Sufficient measurements must be made at the cross-section to determine the degree of

mixing at the prospective site under different flow conditions and to verify that cross-

section variability at the site is not greater than conditions necessary to meet data-

quality objectives. Cross-section measurements must continue to be made after

equipment installation to ensure that the measurements are representative of the

stream during all seasons and hydrographic flow conditions (Wagner et al. 2006).

Additional information on cross-section surveys can be found in Chapter 4.1, In-Situ

Field Meter and Transect Data Collection Protocol (Hoger 2020).

Some aquatic environments may present unique challenges for optimal site location.

Lateral mixing in large rivers often is not complete for tens of miles downstream from a

tributary or outfall. Therefore, a location near the streambank may be more

representative of local runoff or affected by point-source discharges upstream, whereas

a location in the channel center may be more representative of areas farther upstream

in the drainage basin. Turbulent streamflow may aid in mixing, but turbulence can

create problems in monitoring field parameters, such as DO or turbidity. The ideal

location for a CIM site is often one that is best for measuring surface-water discharge.

Both purposes require a representative site that approaches uniform conditions across

the entire width of the stream.

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Instream Site Selection

The measurement point in the vertical dimension also needs to be appropriate for the

primary purpose of the monitoring installation. Flow condition should be considered

during this evaluation. For a medium to small stream with alternating pools and riffles,

the best flow and mixing occurs in the riffle portion of the stream; however, if flooding

changes the locations of shoals upstream of the monitoring site, the measurement point

may no longer represent the overall water quality characteristics of the waterbody.

Streams subject to substantial bed movement can result in the sensors being lost or

located out of water following a major streamflow event, or at a point no longer

representative of the flow. A site may be ideal for monitoring high flow but not

satisfactory during low flows.

Assessment of a site also is dependent on fouling potential, ease of access, and

susceptibility to vandalism. The configuration and placement of water quality monitoring

sensors in cold regions require additional considerations in order to obtain data during

periods of ice formation. Overall, a monitoring site should be safe and accessible, meet

minimum depth requirements of the equipment, avoid vandalism, and be characteristic

of the system. It may be necessary to reconnoiter the site under several flow conditions

before a determination is made. The site should continue to be evaluated throughout

the deployment, and if necessary, the monitor moved to a more appropriate location.

Monitor Selection

According to Wagner et al. (2006) the selection of a water quality monitor involves four

major interrelated elements: (1) the purpose of the data collection, (2) the type of

installation, (3) the type of sensor deployed at the installation, and (4) the specific

sensors needed to satisfy the accuracy and precision requirements of the data-quality

objectives. The DEP Water Quality Division uses a wide variety of instream monitors;

however, these monitors are all designed to work with DEP’s preferred monitor

configuration which eliminates the need to consider the type of installation (element

number 2 above).

Sensors are available as individual instruments or as a single combined instrument that

has several different sensors in various combinations. For clarity, in this protocol, a

sensor is the fixed or detachable part of the instrument that measures a particular field

parameter. A group of sensors configured together commonly is referred to as a sonde.

A sonde typically has a single recording unit or electronic data logger to record the

output of multiple sensors. The term monitor refers to the combination of sensor(s) and

the recording unit or data logger (Wagner et al. 2006). The most widely used water

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quality sensors in monitoring installations are water temperature, conductivity, DO, pH,

turbidity, and water stage. These sensors are the focus of this protocol.

Monitor Configuration

The monitor configuration preferred by DEP is an internal-logging, multi-parameter,

recording monitor that is entirely immersed and requires no external power. Power is

supplied by conventional batteries located internally, and sensor data are stored within

the sonde on nonvolatile recording devices. DEP uses monitors from multiple

companies but prefers models that can provide live readings to facilitate fouling and

calibration checks outlined below, and that allow calibration of the sensors to minimize

corrections of recorded data. Models that do not have these features can be used but

performing the necessary checks will be more challenging.

The standard deployment procedure consists of the monitor placed inside a well vented

schedule 80 Poly Vinyl Chloride (PVC) shroud attached to two form stakes with

carabiners and a steel cable (Figure 1). The form stakes are driven into the sediment

paralleled to stream flow. In this configuration, the stakes act as a primary and

secondary anchor for the monitor. Smaller monitors are also configured in this style,

however shrouds are smaller and may only have one form stake anchor based on

stream flow characteristics (Figure 2). In both small and large monitor configurations,

equipment is covered by large rocks for protection and concealment. Figure 3 illustrates

the standard instream configuration for DEP.

The primary advantages of the internal-logging configuration are that AC power or large

batteries and shelters are not needed. As a result, the upfront cost associated with

emplacement is greatly reduced and mobility of the monitors is increased. Additionally,

mid-channel deployment is much easier. Some disadvantages to this configuration

include the lack of telemetry and solar panels which increases the need for regular

maintenance and data retrieval visits. Additional monitor configurations are discussed in

Wagner et al. (2006).

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Figure 3. The standard instream configuration for monitors operated by DEP. Large, but

manageable, rocks (not shown in figure) are placed around the monitor for stability,

concealment, and protection.

Adjusting for Sediment Fouling

In streams characterized by high sediment deposition, monitors can quickly become

buried if deployed on the stream bottom. Even when monitors are equipped with wipers,

sediment accumulation can quickly lead to erratic readings that usually render data

unusable. In these environments, DEP suggests trying the following adjustments to the

standard deployment:

• Prop the monitor up slightly off the bottom with a small rock. With mild

deposition, this procedure is often sufficient.

• Angle the monitoring into the flow of the stream. Support this angle with the rocks

covering the sonde. This angle can increase flow across the probes and help

flush sediment.

• Shorten the PVC shroud. When the sonde guard and probes protrude from the

PVC, sediment accumulation is often reduced. Employing this strategy does

leave the probes less protected.

When severe sediment fouling conditions are present, it is necessary to deploy the

sonde in a way that keeps it off the stream bottom. One effective strategy is to anchor a

section of PVC to rocks or trees on the bank and slide the sonde down the PVC into the

water (Figure 4). A long bolt through the PVC is used to create a stop for the sonde at

the correct height, suspending it in the water column. Cross-section surveys are crucial

when employing this strategy to ensure that the water quality near the bank is

representative of the stream. Another option to keep the sonde off the bottom of the

stream is to suspend the sonde using a bridge or bridge pier (Figure 5). With either

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work is the Celsius scale. Modern thermistors can measure temperature to ± 0.1°C, but

accuracy should be verified with the manufacturer (Wilde 2006).

Specific Conductance

Electrical conductivity is a measure of the capacity of water to conduct an electrical

current and is a function of the types and quantities of dissolved substances in water.

As concentrations of dissolved ions increase, conductivity also increases. Specific

conductance is the temperature specific calculation of conductivity expressed in units of

microsiemens per centimeter (Radtke et al. 2005). DEP’s Water Quality Division

measures and reports specific conductance in microsiemens per centimeter (μS/cm) at

25 °C. Specific conductance measurements can be a good surrogate for total dissolved

solids and total ion concentrations, but there is no universal linear relation between total

dissolved solids (TDS) and specific conductance. A relation between specific conduc-

tance and constituent concentration must be determined for each site (Radtke et al.

2005). In many circumstances, the amplitude and variability of specific conductance is

not sufficient to predict other constituent concentrations (such as TDS) within the

stream. Therefore, it is important to note that TDS measurements from monitors are

calculated from conductivity and are not always accurate or reliable without further

evaluation of the ion constituents.

Monitoring systems used by DEP usually contain automatic temperature compensation

circuits to compensate specific conductance to 25 °C. This can be verified by checking

the manufacturer’s instruction manual. Most modern sensors are designed to measure

specific conductance in the range of 0–2,000 μS/cm or higher. Specific conductance

sensors are reliable, accurate, and durable but are susceptible to fouling from aquatic

organisms and sediment. Typical accuracy of specific conductance probes in freshwater

is ± 1 μS/cm or 0.5% of reading, whichever is greater.

pH

The pH of a solution is a measure of the effective hydrogen-ion concentration. Solutions

having a pH below 7 are described as acidic, and solutions with a pH greater than 7 are

described as basic or alkaline. Dissolved gases, such as carbon dioxide, hydrogen

sulfide, and ammonia, affect pH. Degasification (for example, loss of carbon dioxide) or

precipitation of a solid phase (for example, calcium carbonate) and other chemical,

physical, and biological reactions may cause the pH of a water sample to change

appreciably soon after sample collection (Ritz and Collins 2008).

The electrometric pH-measurement procedure, using a hydrogen-ion electrode,

commonly is used in continuous water quality pH sensors. A correctly calibrated pH

sensor can accurately measure pH to ± 0.2 pH units; however, the sensor can be

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scratched, broken, or fouled easily. If flow rates are high, the accuracy of the pH

measurement can be affected by streaming-potential effects (Ritz and Collins 2008).

These types of probes typically have a life-span of 1-2 years as the reference solution is

depleted (some manufacturers offer refillable reference solution receptacles to extend

the life of the probe). This life-span is affected by the ionic concentration of the solution

and water in which a probe is stored and deployed. Lower ionic concentrations more

quickly deplete the reference solution. The pH mV readings provide insight to the

current condition of the probe. DEP’s Water Quality Division recommends recording

these values at each calibration. The pH mV values should be within the following

criteria:

• Buffer 7: readings should be -50 to 50 mV, 0 mV is ideal

• Buffer 4: readings should be +165 to +180 mV from the buffer 7 reading

• Buffer 10: readings should be -165 to -180 mV from the buffer 7 reading.

As pH probes age and reach the thresholds listed above, they can become slower to

respond and the data becomes increasingly erratic (Figure 7).

Dissolved Oxygen

Dissolved oxygen of streams is produced by diffusion from atmospheric oxygen and

photosynthetic productivity, and drives chemical reactions within and above the

substrate and is critical for the survival of aquatic organisms. Expected DO

concentrations vary based on temperature and other factors but are generally from 6 to

14 milligrams per liter (mg/L). Dissolved oxygen is also reported as percent saturation,

which will decrease with increasing water temperature, and increase with increasing

atmospheric pressure. Another impact to DO solubility is salinity. Dissolved oxygen

measurements in waters that have specific conductance values >2,000 μS/cm should

be corrected for salinity. Most modern sensors automatically compensate for the effects

of salinity or have manual compensation techniques, but this should be verified by

checking the manufacturer’s instruction manual (Rounds et al. 2013).

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or reducing chemical agent, such as a chemical spill, by metal-rich drainage water, or

by organic-rich waters. Typical accuracy of a membrane-type DO sensor is ± 0.2 mg/L

or 2% of the reading, whichever is greater (within the range 0-20 mg/L).

Turbidity

Turbidity is defined as an expression of the optical properties of a sample that cause

light to be scattered and absorbed rather than transmitted through a sample. Most

turbidity sensors emit light from an LED into the water and measure the light that

scatters or is absorbed by the suspended particles in the water. The sensor response is

related to the wavelength of the incident light and the size, shape, and composition of

the particulate matter in the water. The effect of temperature on turbidity sensors is

minimal, and the software for modern sensors provides temperature compensation.

Sensors that are regularly maintained and calibrated are somewhat error free. Turbidity

sensors should be calibrated directly rather than by comparison with another meter

(Anderson 2005). Turbidity can be reported in several different units depending on what

type of sensor is being used. One common unit of measure is nephelometric turbidity

units (NTU); however, DEP uses sensors that report data in formazin nephelometric

units (FNU), because these sensors use near infra-red light (780-900nm) with a

detection angle of 90°. For the purposes of measuring a calibration solution, turbidity

units are equivalent; however, instruments may not produce equivalent results for

environmental samples, which is why different turbidity units are required. Turbidity

sensors typically have a range of 0–1,000 FNU and an accuracy of ± 0.3 FNU or 2%,

whichever is greater. Some sensors can report values reliably up to about 4,000 FNU.

Water Stage

Most stream monitors employ pressure transducers to measure water stage. The

physical force measured is made up of various factors such as atmospheric pressure,

temperature fluctuations, and water pressure. Pressure transducers convert physical

force into an electronic signal to be calculated into water stage. Most modern stage

sensors will automatically compensate for fluctuations in temperature.

Water stage sensors come in two common forms, non-vented and vented. Non-vented

(absolute) sensors measure both atmospheric pressure and water stage

simultaneously. Resulting data from non-vented sensors require corrections provided by

either a separate barometric pressure monitor or by using barometric data from a near-

by airport/weather station. Vented (gage) sensors employ a vent tube that is open to the

air. As a result, these sensors will automatically compensate for the changes in

atmospheric pressure as long as the vent tube remains unobstructed and dry.

Due to the monitor deployment procedure, DEP generally uses non-vented pressure

sensors to record water stage. A nearby barometric pressure monitor is typically used to

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correct for atmospheric pressure changes. If separate barometric monitors are used,

they should be set to the same sampling interval as the stream monitor to facilitate the

data management process. Both stream monitor and barometric monitor must be

calibrated at the respective monitoring site before logging begins.

When considering the use of a non-vented water stage sensor and a separate

barometric monitor it is important to account for distance (between stream and

barometric monitor) and elevation. DEP’s Water Quality Division generally recommends

that the distance between stream monitor and barometric monitor is not greater than 30

miles. This reduces the potential for barometric correction errors due to localized

weather patterns. As a general rule for Pennsylvania, barometric pressure changes with

elevation at a rate of 1 in-Hg for every 1000 ft. If the elevation difference from stream

monitor to barometric monitor is minimal, then no elevation compensation is needed.

Correcting for barometric pressure using another instrument adds to the total potential

error within the data set. Unless the accuracy of each instrument can be documented,

and verified depth measurements are taken at each maintenance visit, the data must be

dealt with in a qualitative manner.

USE OF FIELD METERS

The three major uses for a field meter during servicing of a continuous water quality

monitor are (1) as a general check of reasonableness of monitor readings, (2) as an

independent check of environmental changes during the service interval, and (3) to

make cross-section surveys or vertical profiles in order to verify the representativeness

of the location of the sonde in the aquatic environment. Data collected by the field meter

should not be used to calibrate the instream monitor. However, some instream monitors

cannot be calibrated, in which case the calibrated field meter must be used in the

calibration correction of monitor records. In addition, DEP may use a calibrated field

meter measurement for undocumented data error corrections (a potential result from the

DEP deployment procedure, discussed in detail later). Independent field measurements

must be made before, during, and after servicing the monitor in order to document

environmental changes during the service. Side-by-side measurements between the

field meter and the monitor are made by placing the calibrated field meter as close to

the monitor sensors as possible. The field meter should refresh data on the display at

10 second intervals, or more frequently, to ensure the most accurate data are recorded

during measurements.

Before site visits, all field meters should be checked for operation and accuracy.

Minimum calibration frequency should be once per day in the field. Recalibration will be

necessary when a sensor is replaced, and during dramatic changes in elevation or

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barometric pressure. Calibrations must be recorded in instrument log sheets. The Field

Meter Calibration Form is provided in Appendix B.1. DEP recommends that all sensors

are calibrated in accordance with manufacturer’s specifications. Additional tips and

guidance on field meter use are provided in Hoger (2020).

MONITOR OPERATION AND MAINTENANCE

The objective during continuous instream monitoring is to obtain the most accurate and

complete record possible. The common operational categories include maintenance

frequency, field visits, troubleshooting, and comprehensive record keeping.

This instream monitoring protocol details one operating procedure designed for well-

mixed, stable, and relativity slow changing systems. Slowly changing is defined as

changes in field measurements during maintenance that are less than the calibration

criteria (see Sensor Field Calibration). The wadeable streams of Pennsylvania generally

fall into this category.

Field Notes and Instrument Logs

Logs and field notes are essential for accurate and efficient record processing.

Operation and maintenance records are maintained in an electronic spreadsheet similar

to the USGS standard field form for water quality monitors in Wagner et al. (2006). A

copy of the spreadsheet is available upon request. The deployment and maintenance

visit field forms from the spreadsheet are provided in Appendices B.5 and B.6,

respectively. Field-note requirements for instream monitors include:

1. Station name

2. Date and time of measurements

3. Name(s) of data collector(s)

4. Serial number of field meters and monitor

5. Lot numbers and expiration of dates of standard solutions

6. Location description and picture of monitor in the stream

7. Name of file downloaded from the monitor

8. Monitor values, field meter values, and corresponding time for cleaning checks

(for fouling), calibration checks, calibrations/recalibrations, and final readings

9. Battery voltage of monitor at departure and if the batteries were replaced, name

of new file, and start time of logging

10. Notes on sensor/monitor changes or replacements, and other comments that

facilitate processing of the record

11. Cross-section survey data (locations of points, measured values, and corresponding times), and monitor values before and after the cross-section survey (Hoger 2020)

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12. Measured flow or gage-height data.

Logging Interval and Settings

Most monitors can be set for a wide range of recording intervals. The interval used

should account for the expected rate of change in the water quality parameters being

measured. In most streams in Pennsylvania, a 30-minute interval adequately captures

changes and therefore is the most common interval used by DEP. “Flashy” streams,

that are high gradient or have a significant amount of impervious cover in the

watershed, change more rapidly and may necessitate a 15-minute interval. When

recording turbidity, many prefer to have the additional data points of a 15-minute interval

so that the curves of storm events are better represented. A five-minute interval has

even been used for a deployment directly downstream of a point-source discharge to

capture its short, concentrated discharges. When deploying the monitor independently,

however, battery life can be an important consideration and may necessitate a longer

interval or a unit with greater battery capacity.

To avoid confusion when comparing continuous data and discrete samples, DEP does

not recognize daylight saving time when working with its continuous data. All sondes

are set to record in EST (UTC-05:00) and all associated measurements, such as

discretes, should be recorded in the same manner to avoid confusion.

Setting the monitor to log is different with each unit, so follow the manufacturer’s

instructions. When setting the monitor to log, make note of the interval, name of the file,

battery life, and start time. If possible, DEP recommends delaying the start time by at

least one hour to allow the monitor plenty of time to stabilize. It is also recommended

that the battery voltage parameter is set to record throughout the deployment to aid in

determination of battery life. Finally, employing a consistent naming convention (e.g.,

Newport_1, Newport_2, etc.) can aid in the organization and uploading of files later.

Maintenance Frequency

DEP Water Quality Division monitors are typically placed at a station for approximately

one year and revisited 10-12 times during the deployment. However, maintenance

frequency will generally depend on fouling rate of the sensors, which varies by sensor

type, environmental conditions, and season. The performance of water temperature,

specific conductance, and water stage sensors tends to be less affected by fouling than

DO, pH, and turbidity sensors. Wiper mechanisms on turbidity and optical DO sensors

can substantially decrease fouling in certain aquatic environments. Monitoring sites with

nutrient-enriched waters and moderate to high temperatures may require more frequent

maintenance. In cases of severe environmental fouling or in remote locations, the use of

an observer (e.g., landowner, volunteer, or local watershed group) to provide more

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frequent maintenance to the water quality monitor should be considered. Monitoring

disruptions as a result of equipment malfunction, sedimentation, electrical disruption,

debris, ice, or vandalism also may require additional site visits.

Field Visits

Field visits are undoubtedly the most important step in certifying that quality data are

being recorded. The purpose of field visits is to verify that a sensor is working

accurately, upload recorded data, confirm calibrations, and provide a reference point for

subsequent data management. Quality assurance is accomplished by recording field

fouling observations before and after cleaning sensor readings in the environment,

calibration checks, and final readings. It is important to conduct field checks at, or close

to, the monitor’s deployment location to record checks that best represent stream

conditions recorded by the monitor. To prevent complications from freezing, caution

should be exercised when performing maintenance at temperatures below 0˚C, and

maintenance should be avoided at temperatures below -7°C or 20˚F.

The standard protocol for servicing instream monitors is described below:

1. Obtain a discrete measurement from a clean, calibrated field meter at the sonde

location

2. Remove sonde from the monitoring location being careful to minimize

disturbance

3. Connect monitor to field instrument (i.e., computer or handheld device)

a. If monitor is to be submerged during read-out, ensure the cable is

designed to operate under water

b. Stop unattened monitoring

c. Upload data

d. Do a quick review of data to detect any data abnormalities or defects in probes

e. Record any significant fouling observed during monitor removal

4. Conduct before-cleaning, initial monitor inspection

a. Record time, readings, and monitor conditions

b. With an independent field meter, record instream readings and time near

the monitor

5. Clean sensors (see Sensor Field Cleaning)

6. Return sonde to the stream and conduct after-cleaning monitor inspection

a. Record monitor readings and time

b. Using an independent field meter, record instream readings near the

monitor

7. Remove sonde, and check calibration (see Sensor Field Calibration)

a. Record calibration-check values

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b. Recalibrate if necessary

8. Conduct final readings

a. Record monitor readings and time

b. Using an independent field meter, observe and record readings near the

monitor

9. Restart unattended sampling with appropriate logging interval, start time, and file

name

a. DEP recommends a delay start time of 1-2 hours to allow equipment to

acclimate following disturbance of the site

b. Check monitor’s battery levels, change if necessary

10. Return sonde to monitoring location and inspect anchoring equipment and

shroud for deterioration and damage.

The initial monitor readings during a field visit may not accurately represent the

recording conditions because of disturbance while retrieving and connecting the

monitor. Care should be taken to minimize disturbance and placement of the monitor

during retrieval, and the field meter should be upstream of any disturbed area. If

observed, significant fouling should be noted in the field sheet. Monitors and field

meters should be given ample time to stabilize before readings are recorded. During

extreme conditions (extreme cold/heat, early morning, etc.) monitors and field meters

may require more time to stabilize.

Comparison of the initial monitor and field meter readings provides a sense of the

reasonableness of the recorded data and an indication of any potential errors. The

disturbance of a stand-alone monitor during retrieval can prevent the capture of a true

fouling error, so care should be taken to minimize this disturbance as much as possible.

The difference between the last recorded value on the deployed monitor and the initial

reading after retrieval helps establish any effects of disturbance during the monitor

retrieval, and can be used to calculate what is then referred to as undocumented fouling

(described in further detail in Data Correction). A monitor that is connected to an

external recording device can typically be accessed for initial side-by-side readings

without disturbance—eliminating the potential for undocumented fouling error.

After initial sensor readings, the monitor’s sensors are inspected for signs of fouling.

These observations are recorded in the field notes before cleaning, and then individual

sensors are cleaned according to the manufacturer’s specifications. The monitor is then

returned to the water. Cleaned sensor readings, field meter readings, and times are

recorded in the field notes. If the conditions are steady state, the field meter readings

should not change substantially during the time that the monitoring sensors are cleaned.

The observed difference between the initial sensor reading and the cleaned-sensor

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reading is a result of fouling (chemical precipitates, stains, siltation, or biological

growths).

After cleaned-sensor readings are recorded, the monitor is removed from the water,

calibration is checked in calibration standard solutions, and the readings are recorded.

Differences between the cleaned-sensor readings in calibration standard solutions and

the expected reading in these solutions are the result of calibration error (drift). The

sonde is recalibrated if necessary, and replaced in the aquatic environment for a final

reading.

The set of final readings may not necessarily accurately represent the start of the new

record period due to site disturbance. Care should be taken to minimize disturbance

and placement of the monitor and field meters should be upstream of any disturbed

area. Monitors and field meters should be given ample time to stabilize before readings

are recorded. Final sensor readings should be used in conjunction with the first data

record recorded to determine any affect disturbance may have had on final sensor

readings, field meter readings, and data collected as part of the new record period.

Sensor Field Cleaning

During the cleaning process, care should be taken to ensure that the electrical

connections are kept clean and dry. Water on the connector pins can cause erratic

readings. For this reason, a container of compressed air is useful. Procedures for

cleaning specific sensors, as described below, are general guidelines and should not

replace manufacturer’s instructions. Most commercial thermistors can be cleaned with a

soft-bristle brush and rinsed with deionized water (Wilde 2004).

Rinse specific conductance sensors thoroughly with de-ionized water before and after

making a measurement. Oily residue or other chemical residues (salts) can be removed

by using a detergent solution. Specific conductance sensors can soak in detergent

solution for many hours without damage. Carbon and stainless-steel sensors can be

cleaned with a soft brush, but platinum-coated sensors should never be cleaned with a

brush (Radtke et al. 2005). Platinum-coated sensors may be cleaned with a cotton

swab.

The pH electrode must be kept clean in order to produce accurate pH values. The body

of the electrode should be thoroughly rinsed with de-ionized water before and after use.

In general, this is the only routine cleaning needed for pH electrodes; however, in cases

of extreme fouling or contamination, the manufacturer’s cleaning instructions must be

followed (Ritz and Collins 2008).

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Optical DO sensors are cleaned with a soft bristle brush and rinsed with deionized

water. If the optical DO sensor is equipped with a wiper, ensure the motor is operating

properly and parking in the correct position. Also, ensure that the wiping mechanism

(pad or brush) is in good condition and clean. The black membrane is sometimes

scratched off by coarse debris caught in the wiper. The membrane should be replaced if

over half of the surface has been worn off.

Routine cleaning of polarographic DO sensors involves using a soft-bristle brush to

remove silt from the outside of the sensor, wiping the membrane with a damp, lint-free

cotton swab (available at local electronics stores), and rinsing with de-ionized water.

The sensor usually is covered with a permeable membrane and filled with a potassium

chloride solution. The membrane is fouled easily and typically will need to be replaced

every 2 to 4 weeks. When the membrane is replaced, the potassium chloride solution

must be rinsed out of the sensor with de-ionized water followed by several rinses with

potassium chloride solution before the sensor is refilled. The membrane must be

replaced with care so that the surface of the membrane is not damaged or

contaminated with grease, and no bubbles are trapped beneath the membrane. The

surface of the membrane should be smooth, and the membrane should be secured

tightly with the retaining ring. The sensor must be stored in water for a minimum of 2 to

4 hours, preferably longer, to relax the membrane before installation and calibration.

Because of the time required to relax the membrane, replacement of a membrane

during a field visit would require having a pre-relaxed membrane on hand to allow for

immediate calibration, otherwise it would be necessary to revisit the site after replacing

the membrane waiting the required amount of time before calibration. The retaining ring

must be replaced annually or more frequently to prevent loss of electrolytes. Replacing

the retaining ring when membranes are changed ensures a tight seal.

The gold cathode of the DO sensor also can be fouled with silver over an extended

period of time, and a special abrasive tool usually is required to recondition the sensor.

A fouled anode, usually indicated by the white silver electrode turning gray or black, can

prevent successful calibration. As with the cathode, the sensor anode usually can be

reconditioned following the manufacturer’s instructions. Following reconditioning, the

sensor cup must be rinsed, refilled with fresh potassium chloride solution, and a new

membrane installed (Rounds et al. 2013).

Turbidity sensors are extremely susceptible to fouling; thus, frequent maintenance trips

may be necessary to prevent fouling of the turbidity sensor in a benthic environment

high in fine sediment, algae accumulation, or other biological or chemical debris. In

environments that cause severe algal fouling, however, algae can accumulate on the

wiper pad preventing complete removal of debris from the optical lens, resulting in

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erratic turbidity data. If the turbidity sensor is not equipped with a mechanical cleaning

device that removes solids accumulation or a shutter that prevents accumulation on the

lens before readings are recorded, reliable data collection is very difficult. Sensors first

should be inspected for damage, ensuring that the optical surfaces of the probe are in

good condition. The wiper pad or other cleaning device should be inspected for wear

and cleaned or replaced if necessary. Before placing the turbidity sensor in standards,

the optic lens should be carefully cleaned with alcohol by using a soft cloth to prevent

scratching (or as recommended by the manufacturer), rinsed three times with turbidity-

free water, and carefully dried. If the readings are unusually high or erratic during the

sensor inspection, entrained air bubbles may be present on the optic lens and must be

removed (Anderson 2005).

Sensor Field Calibration

Calibration should only be performed with standards of known quality. All calibrations

and calibration checks should be completed with at least two standard solutions. At a

minimum, these standard solutions should bracket the entire range of recorded values.

An additional standard may be necessary that is proximate to expected environmental

conditions to insure linearity of sensor response. Frequently used standards are

described for each parameter below.

Field calibration is performed if the cleaned-sensor readings obtained during the

calibration check differ by more than the calibration criteria (Table 1). Spare monitoring

sondes or sensors are used to replace water quality monitors that fail calibration after

troubleshooting steps have been applied (see Troubleshooting Procedures). All

calibration equipment and supplies must be kept clean, stored in protective cases

during transportation, and protected from extreme temperatures.

The following example is helpful for understanding the calibration criteria given in Table

1. The calibration criterion for specific conductance is 5 µS/cm or 3%, whichever is

greater. Therefore, if a sonde is in 1000 µS/cm standard and reported a reading of 1025

µS/cm the reading would still be within the calibration criteria because 3% of 1000

µS/cm (30 µS/cm) is greater than 5 µS/cm. For a calibration check in 100 µS/cm

standard the criteria would be 5 µS/cm, because 5 µS/cm is greater than 3% of 100

µS/cm. Sensors with readings outside of criteria will be recalibrated during the field

visits. Best professional judgment should be used with values that read close to

calibration criteria. Data management is often facilitated by eliminating unnecessary

calibration events.

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Table 1. Calibration criteria for measured parameters.

Field Parameter Calibration Criteria

Temperature ± 0.2°C

Specific Conductance ± 5 µS/cm or 3%, whichever is greater

pH ± 0.2 units

Dissolved Oxygen ± 0.3 mg/L

Turbidity ± 0.5 FNU or ± 5%, whichever is greater

Using Standard Solutions

Expiration dates and lot numbers for the standard solutions must be recorded and

sufficient time provided for the sensor to stabilize in the solution. After the readings in

the calibration standard solutions are checked and recorded (without making any

adjustments), the monitor is recalibrated, if necessary, using the appropriate calibration

standard solutions and following the manufacturer’s calibration procedures. When using

standard solutions, the process for both field checks and calibration are the same. The

sensor, thermistor, and measuring container must be rinsed three times with a standard

solution. Gentle tapping will ensure that no air bubbles are trapped in or on the sensor.

After the series of rinses, fresh standard solution is poured into the calibration cup

ensuring both the thermistor and the sensor are submerged; the sensor values,

calibration standard values, and temperature are recorded in the field log sheet. A

temperature correction may be necessary if the monitor does not have automatic

temperature correction. Standard solution that has been used is discarded. If an error

arises during calibration, see manufacturer’s guidelines or reference Troubleshooting

Procedures below. If these steps fail, the sonde or monitoring sensor must be replaced

and the backup instrument calibrated.

Water Temperature Sensors

Manufacturers generally make no provisions for field calibration of water temperature

sensors, but sensors can be periodically NIST certified to ensure accuracy. The water

temperature sensor and the calibrated field thermistor are placed adjacent to each

other, preferably in flowing water. Sufficient time for temperature equilibration must

elapse before a reading is made. The two water temperature sensors must be read and

recorded simultaneously. If the monitoring water temperature sensor fails to agree

within ± 0.2 °C, troubleshooting steps must be taken; if troubleshooting fails, the sensor

must be replaced, as calibration is not possible.

Specific Conductance Sensors

Specific conductance sensors should be checked with at least two calibration standard

solutions of known quality before any adjustments are made. Most frequently, the two

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checks are 100 µS/cm and 1000 µS/cm, however, if stream conditions are likely to be

greater than 1000 µS/cm, a 5000 µS/cm standard is also used to ensure the measured

values are within the limits of the standards used. In addition, the zero response of the

dry sensor in air should be checked and recorded to ensure linearity of sensor response

at low values. If the sensor-cleaning process fails to bring a specific conductance

sensor within the calibration criteria (Table 1), the sensor must be recalibrated.

Calibration of a specific conductance sensor is a single point calibration. The high

standard should be used for the calibration (often 1000 µS/cm but 5000 µS/cm could be

necessary in some streams) and a lower standard, nearer the ambient conditions (often

100 µS/cm), is used as an intermediate check of linearity after calibration.

pH Sensors

DEP’s Water Quality Division uses three buffers (4, 7, 10) to first check and then, if

necessary, recalibrate pH sensors; this ensures all likely field conditions are bracketed.

It is also necessary during checks and calibration to account for deviations from the

listed pH value of the buffers at different temperatures. The listed value of the buffer is

the pH at a specific temperature (typically 25 C). The manufacturer of the buffer

solution should offer a table of expected values at other temperatures. While these

tables are very similar across major manufacturers, it is important that tables specific to

the manufacturer are used to account for any differences. The tables are often provided

at five-degree increments; however, particularly with the 10 buffer, the five-degree

interval results in significant jumps in the adjusted pH values. To fill these gaps,

regression lines can be fitted to the provided data points. An example table of

temperature adjusted pH values and the steps to create the more specific table values

are provided in Appendix B.7.

Expiration dates of the buffer solutions are checked and recorded. The pH mV readings

in each buffer should also be recorded to track the performance of the sensor (see

Sensor Selection for more information). Spare pH sensors or entire backup monitors are

carried in case replacement of the sensor is required.

Dissolved Oxygen Sensors

Dissolved oxygen in water is related to water temperature, atmospheric pressure, and

salinity. Calibration of optical DO sensors should be checked at 100% saturation and

with a fresh zero-DO solution before any adjustments are made. The manufacturer’s

calibration procedures must be followed closely to achieve a calibrated accuracy of ±

0.1 mg/L concentration of DO (Rounds et al. 2013). Theoretical charts based on water

temperature and barometric pressure should be used to confirm calibration of monitors.

A general 100% saturation chart for fresh water is provided in Appendix B.8.

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Customized charts can also be made from the following USGS website:

http://water.usgs.gov/software/DOTABLES/.

Most optical DO sensors can be calibrated with only a one-point calibration, usually at

100-percent saturation, although some sondes have the capability of a two-point

calibration, at zero-percent and 100-percent saturation. For the sondes that are

calibrated only at 100-percent saturation, the DO sensor response is still checked in a

zero-DO sodium sulfite/cobalt chloride solution. A fresh zero-DO standard solution

should be prepared before each monitor visit by dissolving 0.5 gram of sodium sulfite

and 6 – 10 crystals of cobalt chloride in 500 ml of deionized water. The maintainer

should observe the response of the sensor in the solution and remove the sensor at 0.3

mg/L because letting the sensor go completely to zero may cause damage.

Optical DO sensors are calibrated by the manufacturer, and some manuals indicate that

calibration may not be required for up to a year. When calibrated, the user should follow

the manufacturer’s guidance. Regardless of the manufacturer’s claims, the user must

verify the correct operation of the sensor in the local measurement environment. The

standard protocol for servicing should be used for optical DO sensors to quantify the

effects of fouling and calibration drift (Rounds et al. 2013). Recalibration should not be

necessary if calibration checks show the sensor to be in agreement with the calibration

criteria (Table 1) and theoretical chart values. Spare membranes and probes should be

available in case replacement is necessary.

Calibration of polarographic sensors in the field presents a problem because

replacement of the TeflonTM membrane may be required frequently, and the replaced

membrane must be allowed to “relax” in water for 2−4 hours before calibration. One

solution to this problem is to carry into the field clean and serviced spare DO sensors,

stored in water (or moist, saturated air). The replacement DO sensors then can be

calibrated in the field, thus avoiding an interruption in the record and a return site visit

(Rounds et al. 2013).

Turbidity Sensors

Optimal calibrations and checks are made with three standard solutions that cover the

expected range of values, although many sensors only allow for a two-point calibration.

Calibration of the turbidity sensor is made by using standard turbidity solutions (DEP

typically uses 0, 124, and 1010 FNU) and by following the manufacturer’s calibration

instructions. When units only allow a two-point calibration, the zero and high value

solutions (e.g., 1010 FNU) should be used to calibrate, and the middle value (e.g., 124

FNU) solution should be used as a check of linearity after calibration. Deionized or

ultrapure (filtered and deionized) water may be used as a zero turbidity standard if its

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turbidity is measured by a laboratory. Laboratory measurements are necessary because

deionized or ultrapure turbidity can measure as high as 1.0 FNU. The measured

turbidity from the laboratory can be set as an offset calibration in the monitor, or the

data can be offset corrected later in data management. Checking or calibrating the

turbidity sensor should occur in a stable environment with minimal movement, wind, or

exposure to sunlight. Sufficient space between the sensor and the bottom of the

calibration cup is also critical during checks and calibration. Calibration in zero standard

is best performed with the sensor guard in place to simulate conditions when deployed.

Bubbles in the solution can interfere with readings so care should be taken to minimize

bubbles when pouring. Running the wiper before each reading or calibration in each

solution will help remove any remaining bubbles.

When using polymer-based turbidity standards, it is important to understand that

different sensors often read the solution differently. Even different model sensors from

the same manufacturer will likely not read a given turbidity standard the same.

Therefore, only polymer-based solutions with documentation for the specific sensor

should be used. In most cases this means that genuine manufacturer solution is

necessary, as standards from other manufacturers lack the necessary documentation –

making proper calibration impossible. Currently, the only alternative is Formazin

turbidity standard. Formazin turbidity standard has the benefit of being read the same

by all sensors, but it is considered moderately toxic, contains a known carcinogen, and

is more difficult to work. Therefore, DEP recommends using polymer-based turbidity

standard—with the proper, sensor-specific documentation—whenever possible.

Water Stage Sensors

Water stage calibration procedures vary slightly with manufacturer, so DEP

recommends following manufacture’s recommendations. When calibrating water stage

for deployment, it is important to calibrate at the monitoring location. Most non-vented

sensors will calibrate to a depth of zero in the air. This accounts for the elevation and

atmospheric pressure at the monitoring site. Since there is no change in elevation

during deployment and most pressure transducers automatically compensate for water

temperature, the only corrections needed are changes in barometric pressure (see

Calculated Derived Series).

In locations where an immobile reference cannot be found, a staff plate or form/rebar

stake may be employed (Figure 8). A staff plate or stake that is driven into the stream

bed provides a point of reference for consistent manual stage measurements which can

be compared to monitor recordings at the time of a field visit. Staff plate or stake

measurements may serve as verified discrete data for subsequent data management

purposes.

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Table 2. Troubleshooting procedures for common issues encountered in the field.

Symptom Possible Issue Likely Solution

Water Temperature

Thermistor is not reading

accurately

Dirty sensor Clean sensor

Erratic readings Poor or wet connection Dry and tighten connection

Slow to stabilize Dirty sensor Clean sensor

Reading off scale Failure in electronics Replace sensor or monitor


Will not calibrate Standard solution old or

contaminated

Use fresh standard solution

Electrodes dirty Clean with brush

Air trapped around sensor Tap gently to expel air

Weak batteries Replace batteries


Requires frequent

calibration

Broken cables Replace cables

Replace monitor

pH

Will not calibrate Standard solution old or

contaminated

Use fresh standard solution

Faulty Sensor Replace sensor


Defective sensor Replace sensor

Slow response time Dirty sensor bulb Clean sensor

Water is cold or of low ionic strength Be patient

Dissolved Oxygen

Will not calibrate Membrane damaged Replace membrane

Electrolyte diluted Replace membrane and electrolyte


Fouled sensor Check for obstructions or replace

Slow to stabilize Gold cathode tarnished Buff with eraser or recondition sensor

Fouled membrane Replace membrane and recondition

Silver anode corroded Replace sensor and soak fouled

sensor in 3% ammonia for 24 hours

Will not zero Zero DO solution contains oxygen Add additional sodium

Zero DO solution is old Mix fresh solution

Turbidity

Unusually high or erratic

readings

Entrained air bubbles on the sensor Gently tap or run wipe cycle

Damaged Sensor Replace sensor

Dirty sensor Clean, using soft brush

Poor or wet connection Dry and tighten connection

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DATA MANAGEMENT

The data management process verifies data, corrects for errors, and sets a grade based

on overall quality of the record. Accurate and concise field notes and logs are

indispensable in processing collected data. The steps in processing data records are:

initial data evaluation, data import, data corrections, grading, and final approval. Data

Management should begin when the record is uploaded in the field to ensure the

equipment is operating properly. Data management is not complete until the monitor is

pulled from the stream, data is graded and approved, and a final Continuous Instream

Monitoring Report (CIMR) is written, reviewed, and approved by DEP staff. DEP uses

Aquarius software (current version: 3.10) from Aquatic Informatics for all data

management. This protocol will cover basic operation of the Aquarius software for data

management purposes. Two interfaces in Aquarius software aid in data management,

Whiteboard and Springboard. This protocol focuses on data management within the

Aquarius Springboard application, a more user-friendly environment than Aquarius

Whiteboard. Additional training and support can be found at:

http://www.aquaticinformatics.com/support.

Initial Data Evaluation

The initial data evaluation is conducted both in the field and back in the office to verify

the accuracy of data transfer to the database, to identify erroneous data, and to confirm

the monitor is operating as expected. Once data are uploaded from the monitor, they

should be quickly reviewed to identify any inconsistencies. Ideally the manufacturer’s

software will allow the data to be displayed visually. This will allow for a quick

determination of the reasonableness of the data including expected range, identification

of erratic readings, and/or presence of expected behavior like diel swings or response to

changes in flow. Various formats are available to store raw field data. Data formatting

will typically depend on the make of the monitor used.

Data Import

Creating New Stations

Data are organized in a cascading folder structure. Stations are organized by internal

(to DEP) or external data, basin, HUC, stream name, and finally a specific location.

Stations are considered unique if they could represent different water quality conditions.

The two main considerations are cross-sectional variability and distance upstream or

downstream. If cross-sectional surveys demonstrate variations in water quality, DEP

may deploy monitors in multiple locations horizontally across the stream; In Aquarius,

each are considered unique stations. Deployments >100 m apart upstream or

downstream are considered unique stations, however, deployments separated by

shorter distances may be considered unique if conditions warrant. Most deployments by

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DEP are in relatively shallow, lotic systems; therefore, vertical variations within the

water column are often not an issue. If introduced, this variable should also be

considered in creating a new station.

Before data can be uploaded to the database server, a station with the appropriate time

series needs to be created. To create a new station first navigate to the correct stream

in the folder structure. If the stream or HUC folders are missing, they first need to be

created. To create the location either right-click on the stream folder and select “New

Location” or highlight the stream folder and click the Location Manager button on the

toolbar. Under the General tab, the following fields need to be populated:

o Identifier: This is a unique number consisting of the ComID from the

NHDFlowline the station is on, followed by a unique identifier. If the ComID of the

NHDFlowline was 57465467 the Identifier of the first site on that segment would

be 57465467-001. If another station is located on that segment at the same time

or in the future, its Identifier would be 57465467-002, etc.

o Name: The name should include both the stream name and a short description

of the location such as a road crossing (e.g., Little Beaver Creek at Little Beaver

Rd). Including the stream name allows for better search results when using

Aquarius. The short location description should be descriptive enough that

anyone can understand its general location.

o Type: Water Quality Site

o Description: Additional information can be added here such as a more thorough

description of the location, whether the location is part of a specific study, etc.

o Latitude: Decimal degree format

o Longitude: Decimal degree format

o Elevation: Required in Aquarius when using latitude/longitude

o Time Zone: DEP does not recognize daylight saving time when working with its

continuous data; therefore, this should be set to EST (UTC-05:00) for all internal

data.

Under the Data Sets tab, create a “Time Series – Basic” for each continuous parameter

to be stored in the database. Field Visit (discrete) Time Series do not need to be

created in advance; they will be auto-generated when data are input. For each basic

time series created, the following fields need to be populated (description and comment

fields are optional):

• Identifier: This field is auto-populated using the location identifier created in the

general tab, the label name created next, and the parameter selected.

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• Label: Short description of the site (often just the stream name or the stream

name and crossing or nearby town). No spaces, capitalize first letter of each

word.

• Parameter: Use the “…” button to search for a parameter if not in the recently

used parameters in the drop-down box. For temperature, use “Water

Temperature” not “Temperature”. There are many options for turbidity, “Water

Turbidity” is the standard parameter used by DEP. Depth should be entered

under “Stage”.

• Units: Select the appropriate units.

• Time Zone: EST (UTC-05:00).

• Gap Tolerance: If time between readings is greater than the value set in this

box, Aquarius implies a gap and the points are not connected. Set this value to

one minute greater than the recording interval (e.g., if interval is 30 minutes Gap

Tolerance should be 31).

Uploading Data

Data can be brought into Aquarius from multiple formats. The most common file type is

a delimited text file such as “.csv”; however, Aquarius will recognize some

manufacturer-specific files (e.g., YSI’s “.dat” file). To upload data, navigate to the site

location in Springboard and then select the “Append to Logger” icon . This tool allows

for the manual uploading of data and the setup of hot folders, an automated upload

process. In the Append to Logger toolbox, first select the file for upload. The second

step is to tell Aquarius how to interpret the file. If a configuration file has already been

created, select it and click the “Load File” button. (Note that configuration files can be

used for different sites as long as the formatting of the files is identical [i.e., headers,

parameters, parameter order, etc.]) If a configuration file has not been created, it will be

necessary to use the configuration toolbox by clicking “Config Settings…”.

Config File Setup

1. Select file type: will most likely be “Text File (CSV, etc)”. Click Next.

2. Make sure the proper delimiter is selected and establish the row to start import

and number of headers. The number of headers used is not important but it is

helpful to include enough to see what the parameter is for the next step. If done

correctly, data will be broken into appropriate columns, header rows will be gray,

and data rows will be white (Figure 9). Click Next.

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To add additional file type parsers within the hot folder, go through the same process in

the Append toolbox as above. Once the parameter has been designated for a specific

parameter, an Add to Hot Folder button will appear. This process will be identical to the

first time around, just specify a description and a filename filter. To change any of these

specifications later, go through the same process in the Append toolbox and select the

Manage Hot Folder button that will appear.

Important Notes Regarding Hot Folders

• Consistency of the naming of files and the layout of the content within the files

themselves is imperative; therefore:

o Give files a consistent name for each site (e.g., Newport1, Newport2, etc.)

o Do not edit the content of files before uploading. While technically this is

not an issue, the edited files are more likely to be inconsistent than the

original, equipment-generated file. Therefore, do not edit or clean up

headers, move columns, etc.

o Do not edit the parameters during deployment. This means do not add or

delete parameters or in the case of Eureka sondes, do not change the

order of display. This would in turn change the layout of the data file, so

the Config File will need updated.

o If the recording sonde is changed, it may be necessary to update the

Config File unless the data files are formatted in the same way.

• The Config File that is associated with the hot folder is copied and stored

elsewhere once the association is made. This means that after making any edits

to a Config File in the Append window, the file must be reassociated to the hot

folder.

Entering Discrete Data

Discrete data are stored as Field Visits in Aquarius. With the site selected, click the

Field Visit icon . Select New from the toolbar, and choose Field Visit. Enter the date

and time of the discrete reading. Make sure the new Field Visit is highlighted in the left

column, select New from the toolbar again, and choose Measurement Activity. Click the

green circle with a white plus sign to add a parameter. In the Parameter column, select

a recent parameter from the drop-down menu by clicking the magnifying glass to find a

parameter. Make sure to choose the exact parameter as established for the continuous

data sets. Enter the reading in the Value column and confirm the units in the next

column are correct. Continue adding parameters until all discrete data are entered

(Figure 13). After entering the last value, be sure to click outside the box before clicking

Save.

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Figure 15. Elimination of changes in atmospheric pressure from non-vented stage data

from Skippack Creek at Mainland Rd, PA. Large flow events (e.g., 8/13/13) are

apparent in the raw data, but many small flow events are concealed by changes in

atmospheric pressure.

Table 3. Common barometric units with water column conversion in feet.

Barometric Unit Water Column Conversion (ft)

1 psi 2.3108

1 atm 33.959

1 kPa 0.3352

1 mmHg 0.04469

1 inHg 1.1330

1 mb (hPa) 0.03352

Non-vented stage can be corrected for changes in atmospheric pressure using equation

(1) in an Aquarius calculated-derived series.

(1) y=x1-((x2-a)*b)

Where

x1 = stage (feet) time series

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x2 = atmospheric pressure time series

a = first reading from the atmospheric pressure data file (only subtract the

changes in atmospheric pressure)

b = conversion factor from Table 3 for atmospheric pressure series x2

Other DEP applications of the calculated-derived series have included conversion of

conductivity data to specific conductance and the generation of continuous ammonia

criteria using the acute and chronic formulas from Chapter 93 water quality criteria.

Conductivity can be converted to specific conductance at 25C using equation (2),

assuming a temperature coefficient of 0.0191. This is the standard coefficient used by

YSI water quality instruments.

(2) y=x1/(1+(0.0191*(x2-25)))

Where

x1 = conductivity time series

x2 = temperature time series (C)

Formulas for ammonia criteria from PA §93.7(a), Table 3 have been converted into

equations for acute (3) and chronic (4) criteria time series. These time series are

generated to establish continuous ammonia criteria thresholds and identify critical time

periods. This analysis saves time and money by focusing collection of ammonia grab

samples during these critical periods.

(3) if (x2<10) {

y=0.12*(((1+pow(10,(9.73-x1)))/(1+pow(10,((0.09+(2730/(x2+

273.2)))-x1))))/(1+pow(10,(1.03*(7.32-x1)))))*((pow(10,((0.09+

(2730/(x2+273.2)))-x1)))+1);

}

else {

y=0.12*(1/(1+pow(10,(1.03*(7.32-x1)))))*((pow(10,((0.09+(2730/

(x2+273.2)))-x1)))+1);

}

(4) if (x2<10) {

if (x1<7.7) {y=0.025*(((1+pow(10,(9.73-x1)))/(1+pow(10,((0.09+

(2730/(x2+273.2)))-x1))))/(pow(10,(0.74*(7.7-x1)))))*((pow(10,

((0.09+(2730/(x2+273.2)))-x1)))+1);

}

else {

y=0.025*(((1+pow(10,(9.73-x1)))/(1+pow(10,((0.09+(2730/(x2+

4-67

273.2)))-x1))))/1)*((pow(10,((0.09+(2730/(x2+273.2)))-x1)))+1);

}

}

else {

if (x1<7.7) {

y=0.025*(1/(pow(10,(0.74*(7.7-x1)))))*((pow(10,((0.09+(2730/(x2+

273.2)))-x1)))+1);

}

else {

y=0.025*((pow(10,((0.09+(2730/(x2+273.2)))-x1)))+1);

}

}

Where

x1 = pH time series

x2 = temperature time series (C)

DATA EVALUATION

Systematic adoption of a standardized data-evaluation process, including rating criteria

and maximum allowable limits are vital in finalizing instream monitoring records.

Corrections are completed and then data are graded by the individual responsible for

the data. If the measured error deviates by more than the maximum allowable limits, the

data are deleted and not included in reports or analyses. Final approval of the data is

completed by a separate individual before it can be finalized in a report.

Data Correction

During the deployment period, the sensors can experience calibration drift or be

influenced by various types of fouling (biological growth, accumulation of sediment or

other debris, etc). The checks performed during the field visits allow these influences on

the data to be corrected in post-processing. These checks, however, should not be

blindly applied as corrections without review. If for instance, a correction would move

the continuous data away from both a discrete field measurement and the subsequent

continuous data set, the field check directing that correction is likely erroneous and the

correction should not be applied. Final correction of data is determined by best

professional judgment, guided by the field checks.

Typically, a data correction period begins and ends on servicing dates. If fouling,

calibration, or undocumented errors are small (within the data correction criteria, Table

4), corrections are not necessary. If an error is larger than the criterion, a correction

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should be applied. The decision to apply data corrections when total errors are smaller

than the correction criterion or to correct for other factors is left to best professional

judgment. Corrections to be applied are calculated using the DEP digital CIM Field

Form spreadsheet. The spreadsheet contains correction worksheets for each field

parameter. These worksheets use the field checks performed at the various site visits to

calculate the corrections and errors for each period. These calculations are then used in

Aquarius to apply corrections. If access to the digital field form is not available, manual

error calculations are provided below.

Table 4. Correction criteria for measured field parameters.

Field Parameter Data Correction Criteria


Specific Conductance ± 5 µS/cm or 3%, whichever is greater

pH ± 0.2 units


Turbidity ± 0.5 FNU or ± 5%, whichever is greater

Manual Error Calculations

Quantifying error is performed in accordance with Wagner et al. (2006). Numerically,

total error (Et), as defined by USGS, is equal to the sum of the absolute values of fouling

error (Ef) and calibration drift error (Ec):

(5) Et = |Ef| + |Ec|

When highly reliable discrete data is available, it may be used to bracket the time series

of data for which error is being determined, thereby allowing the computation of an

undocumented error and/or discrete error. This error is equal to the difference between

data corrected for fouling and calibration drift and the discrete data. If discrete data is in

line with the subsequent data set, an undocumented drift correction may be warranted

and the error is referred to as an undocumented error (Eu). If an undocumented drift

correction is not warranted, the difference between the corrected data and the discrete

data is referred to as a discrete error (Ed). If present, these errors should be included in

the total error calculation, modifying equation (5) to:

(6) Et = |Ef| + |Ec| + |Eu| + |Ed|

Total error, as shown in equation (6) may then be applied in data grading.

One important distinction when calculating errors is the difference between a percent

correction and percent error. A percent correction is the calculated percentage that is

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used to correct the continuous data. Percent error is the percentage difference that the

raw continuous data is from the correct value. The difference in the calculations is the

denominator. Correction calculations are made in relation to the original, raw data and

therefore the raw value is used in the denominator. Error calculations are made in

relation to the “true” value and therefore the denominator is the “true” value (e.g.,

reading in standard or buffer, reading after cleaning, etc.). Percent correction should

only be used for correcting data and should not be used as a measure of error. Percent

error should be used to quantify error but should not be used to correct data. When the

unit error is small, the difference between these two percentages will be minimal;

however, as the error increases, the difference between these calculations will quickly

become significant.

Fouling error (Ef), percent fouling correction (%Cf), and percent fouling error (%Ef) may

be determined using equations (7), (8), and (9), as shown below.

(7) Ef = (Sondeac – Sondebc) – (FMac – FMbc)

(8) %Cf = 100 * (Ef / Sondebc)

(9) %Ef = 100 * (Ef / Sondeac)

Where

Sondeac = sonde measurement after cleaning

Sondebc = sonde measurement before cleaning

FMac = field meter measurement after cleaning

FMbc = field meter measurement before cleaning

Calibration drift error (Ec), percent calibration drift correction (%Cc), and percent

calibration drift error (%Ec) may be determined using equations (10), (11) and (12), as

shown below.

(10) Ec = Vstd – Vsonde

(11) %Cc = 100 * [(Vstd – Vsonde) / Vsonde]

(12) %Ec = 100 * [(Vstd – Vsonde) / Vstd]

Where

Vstd = value of the standard or buffer

Vsonde = value measured by the sonde

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Types of Data Corrections

The parameter type and the data obtained during the checks determine the type of

correction applied. Multipoint corrections that bracket the range of recorded values are

the most accurate corrections and should be used whenever possible. Fouling checks

only have a single point of reference (the stream conditions at the time) and therefore,

multipoint corrections cannot be used. Calibration checks for specific conductance, pH,

and turbidity should all consist of multiple standards that bracket the range of values so

those checks should be used to apply a multipoint correction. If a multipoint correction is

not possible, a drift (unit based) or percent correction should be applied. Guidelines for

which correction to use are summarized in Table 5.

In general, percent corrections are preferred over drift corrections because they adjust

to the scale of the measurements like the sensors (a 10 µS/cm difference in specific

conductance at 1000 µS/cm is not equivalent to a 10 µS/cm difference at 100 µS/cm).

Drift corrections are appropriate for temperature because the range is typically small. If

the range is large or is close to zero, a percent correction may be more appropriate.

Drift corrections are also used for pH because it is a log-based scale.

Because of the potential for a very large range of values, turbidity can present some

difficulty in selecting between drift and percent corrections. Because turbidity is often

low during base flow in Pennsylvania streams, a drift correction is usually most

appropriate. Small errors can represent large percent differences when values are low.

Using a percent correction in these circumstances could lead to a dramatic,

inappropriate adjustment to peaks during the period. Likewise, if a maintenance visit

occurred during one of these peak events, a drift correction could lead to large,

inappropriate shifts in the baseline values. Best professional judgment will need to be

used in these circumstances.

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Table 5. Types of corrections to use based on parameter and type of error being

corrected. Undocumented errors should use the same correction type as the fouling

correction listed.

Parameter Correction Type

Fouling Calibration

Temperature Drift* N/A

Specific Conductance Percent Multi-point

pH Drift Multi-point

Dissolved Oxygen Percent Percent

Turbidity Drift** Multi-point

*Percent correction may be appropriate if range is large or values near zero

**Percent correction may be appropriate if range is low

All of the above correction types apply the correction in an incremental, linear fashion

where no correction is applied at the beginning of the period and the correction

increases evenly to the end of the period where the full correction is applied. A less

frequently used correction is an offset correction, which applies the same correction to

all points in the period. One common use for an offset correction is for stage. When a

sonde is returned to the stream, the depth is not always exactly the same as the

previous deployment. This change in stage is consistent for the duration of the

deployment so an offset correction is used to adjust the data to the previous

deployment.

Applying Corrections in Aquarius

Corrections can be applied using alternative procedures; however, using software like

Aquarius has multiple advantages. First, is the tracking of all changes to data. All events

(corrections, deletions, approvals, etc.) are time stamped and logged with the user ID

who made the change. This provides transparency and the ability to implement quality

assurance checks. The second benefit is that all changes are input as layers over the

raw data. These layers provide great flexibility as they can be turned on/off or adjusted

at a later time. It also maintains both raw and corrected datasets without requiring a

complete second set of data to be stored. A third benefit is the easy application of

complex corrections like multipoint corrections. Multipoint corrections apply the

correction not only incrementally and linearly over the period but with respect to the raw

value’s position relative to the multiple checks entered in the system.

To apply corrections in Aquarius, select a parameter and click Data Correction .

Multiple parameters can be selected by holding Ctrl. Select the parameter to edit first;

all other parameters will be considered surrogates and will appear as reference(s) only.

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corrections can be entered in the same correction. Figure 19 shows an example of a 0.2

unit drift correction for pH fouling drift.

Figure 19. Continuous pH data from the Susquehanna River at Rockville, PA with a drift

correction of 0.2 units applied.

Percent Corrections

Percent corrections function similarly to drift corrections but, instead of a unit-based

correction, the correction is a percentage of the raw value. This has most importance

when the range of values is high. Figure 20 shows specific conductance from Goose

Creek at Mosteller Park, PA. This watershed has high impervious cover and the stream

is very “flashy”. Baseline specific conductance is over 800 µS/cm but during rain events

the specific conductance drops dramatically. If a drift correction was applied, corrections

during periods following rain events would be almost four times greater relative to their

raw value; therefore, a percent correction should be used instead.

One difference in the application of the correction in Aquarius is that there is not an

option to enter fouling and calibration corrections in separate boxes for the same

correction. The corrections must be done separately or their sum can be entered and

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no influence on the data because all values are between 7 and 10 pH. The correction is

still entered, however, in order to catalog a record of corrections in Aquarius.

Additionally, if a subsequent correction were to move any data below 7 pH, the 4 buffer

correction would become applicable. Like the percent correction, both the start and end

of the period can be adjusted.

Figure 21. Continuous pH data from the Susquehanna River at Rockville, PA with a

USGS multipoint correction applied. Buffers 4, 7, and 10 were used to check pH

calibration. Values in buffer are entered under Input (3.90, 6.95, 10.20, respectively).

The second column is the correction necessary (0.10, 0.05, -0.20, respectively).

Offset Corrections

Offset corrections are a unit-based correction; however, unlike drift corrections, the

value of adjustment is applied equally to the marked region. Offset corrections can be

used on data that has not been calibrated or was incorrectly calibrated; however,

percent or multipoint corrections can also be used in these circumstances, and often are

the more appropriate procedure. A common use of an offset correction is to correct

stage data when the equipment was returned to the stream at a different depth. This

change in depth is constant for the duration of the next period and is best corrected by

an offset correction (Figure 22).

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to be closely reviewed—and perhaps deleted—if temperature data are found to contain

significant error. Wagner et al. (2006) states that for each 1 C change in temperature,

specific conductance and DO concentrations vary 3 percent. For Yellow Spring

Instrument (YSI) equipment, a 1 C change in temperature affects conductivity, pH, and

DO by 4.0%, 0.5%, and 2.0%, respectively, while a 5 C change in temperature results

in changes of 22.0%, 2.0%, and 9.0%, respectively (YSI Tech Support, personal

communication, January 18, 2017). The equations establishing these relationships are

often proprietary and therefore the exact influence of errors in temperature data cannot

be derived. In addition, without these equations, specific conductance, pH, and DO

cannot be corrected for errors in temperature data.

Undocumented corrections are corrections that move the continuous data based on a

verified discrete sample and/or an adjoining dataset (Figure 23). The type of correction

used depends on the parameter, and should be the same as that used for fouling

corrections. For example, if based on the parameter type the type of fouling correction is

a percent correction, an undocumented correction would also be applied as a percent

correction. It’s important to note that undocumented corrections should be made with

care; the reviewer should ensure that the discrete datum agrees with the continuous

data after redeployment before continuous data are corrected to the discrete.

Grading

Grades are determined based on total error (Et) as defined in equation (2), and are

applied using the Data Correction window in Aquarius. Division of Water Quality

uses a standardized procedure of grading to remain consistent throughout the data

evaluation process (Figure 24). Data are generally classified as either usable or

unusable. Data that fall outside the maximum allowable limits (Table 6) are graded

Unusable and deleted in Aquarius. The maximum allowable limits are established at

approximately 6–10 times the calibration criteria for all standard continuous-monitoring

data-collection activities. These limits and rating criteria are taken from the USGS and

will be considered minimum standards for DEP. The criteria in Table 7 are used to

establish ratings for each period of usable data. The Field Form spreadsheet will help

determine cutoff points between ratings. This process is described in the Correction and

Grading Example.

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Using the last data point in the period usually provides the maximum error applied,

however, if there is a significant change in the data near the end of the period the

maximum error may be at the preceding peak or valley in the data.

When both percent and unit-based thresholds are present in Table 7 (i.e., DO and

turbidity), the procedure to be used is determined in the same way as the calibration

criteria described earlier—whichever procedure would result in a better grade is the

procedure used. This is because when values are low, even small changes can result in

very high percentage differences that would lead to unusable data. Similarly, when

values are high, small percent differences represent large unit differences that would

lead to unusable data.

Selecting whether to use percent or unit-based rating criteria can be problematic with

turbidity data when multiple discretes were taken at drastically different values. Unit-

based ratings are usually used for turbidity data from Pennsylvania streams because

turbidity at base flow is typically low. Discrete readings, however, are often targeted

during peak flow events to facilitate the building of rating curves. These turbidity

discretes at peak flow may have low percent error but high unit-based error. Using

exclusively percent or unit-based rating criteria in this situation will often result in data

being rated as unusable. In order to properly rate the data under these circumstances, it

is necessary to convert the percent error for the discretes taken at peak flows to a unit-

based error that can be summed with the rest of the equation. The turbidity rating scale

has a 1:10 ratio of FNU to percent units and therefore the conversion can be made by

dividing the percent error by 10 to get the unit-based rating equivalent. Alternatively, a

unit-based error can be multiplied by 10 to get a percent rating equivalent.

After the rating is determined based on Et, the grader must determine if any “grade

drops” are warranted. A grade is decreased if there is added uncertainty to the data.

The most common reason for decreasing a grade is a missing verification (fouling

check, calibration check, or discrete measurement during the period). Because the

verification is missing, the grade is dropped by one rating. If two verifications are

missing, the grade should be dropped by two ratings. If all verifications are missing, the

data should be graded unverified. Grade drop determinations are outlined in Figure 24.

To aid the approval process, the person applying the grade should include a short note

in the comment section in Aquarius. The comment section is provided after clicking

Apply. The note, at a minimum, should include the errors calculated and note the

reason for any grade drops.

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Correction and Grading Example

An example of the entire correction and grading process is outlined in Figures 25 and

26 for specific conductance data from the Susquehanna River at Rockville for the period

from 6/9/15 to 7/9/15. The basis for these corrections is taken from the Specific

Conductance tab of the DEP Field Form. These parameter-specific tabs calculate the

errors and anticipated corrections to be applied using the data entered during the

maintenance visit. The section of the Specific Conductance tab related to the period

being corrected is displayed in the upper portion of Figure 25. Many of the cells in this

spreadsheet are populated straight from the individual field visit tabs. The cells

highlighted in purple are the cells to be used to enter corrections in Aquarius or used in

the grading process. Only the cells highlighted in yellow should be edited.

The period was first reviewed visually for outliers or abnormalities such as sections of

erratic readings that needed to be deleted. While these data were slightly more erratic

point-to-point than average, no points needed to be deleted. The erratic behavior of the

sensor was likely due to fouling as the subsequent dataset, after cleaning the sensor,

looked much smoother.

Next, the information from the calibration checks was used to apply a USGS Multi-point

Correction. Because the meter was not calibrated during the maintenance visit on

6/9/15, the correction included adjustments to both the beginning and end of the period.

Under Start Point, the inputs were 0, 103, and 1019 with corrections of 0, -3, and -19,

respectively. End Point inputs were 0, 103, and 1020 with corrections of 0, -3, and -20,

respectively. The Start Point inputs and corrections are the End Point inputs and

corrections from the previous section of the spreadsheet (not depicted in Figure 25).

Once the correction is applied (Figure 26), the error can be calculated. The raw value

was 234.0 µS/cm and the corrected value was 228.6 µS/cm; therefore, using equation

(13) the calibration error for the period is -2.36%:

%Em = 100 * [(228.6 – 234.0) / 228.6] = -2.36%.

It is recommended that calibration corrections are applied first for parameters that

require multi-point corrections. This isolates the multi-point correction, allowing the

above calculation to be made. If the multi-point correction is made after other

corrections, those other corrections will need to be temporarily turned off in Aquarius in

order to make the calibration error calculation.

A Percent Correction was then applied to the period to correct for fouling drift. The Start

Percentage was 0% because the sonde was cleaned at the 6/9/15 maintenance visit,

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and therefore it is assumed that no fouling is present. The End Percentage was -0.88%

which was taken from the Fouling Correction box in the spreadsheet (Figure 25).

If the either correction did not make sense (i.e., data were moved away from the

discrete and/or adjoining datasets), the beginning, end, or both parts of the correction(s)

could have been removed. In this case, the calibration and fouling corrections made

sense because both moved the data closer to the discrete and adjoining datasets;

therefore, both corrections remained applied.

After applying calibration and fouling corrections, the discrete was still 2.0% different

than the corrected data (Figure 26). Because the discrete lined up with the subsequent

dataset, an undocumented correction was deemed appropriate. A Percent Correction

with a Start Percentage of 0.0% and an End Percentage of 2.0% was applied to offset

the undocumented error. It is important to note that this determination was made after

applying the calibration correction to the next period. Because the sensor was calibrated

during the 7/9/15 visit, the beginning of the next period also needed to be corrected.

After all corrections were applied, total error (Et) was calculated using equation (6):

Et = |Ef| + |Ec| + |Eu| + |Ed| = |-0.88%| + |-2.36%| + |-2.0%| + |0.0%| = 5.24%

Fouling error (Ef) was taken from the Fouling Error box of the spreadsheet. This number

happened to be the same as the fouling correction applied because the error was small

in this case. If the error was larger, these numbers would have been different from each

other; therefore, care should be taken to use the correct value. Calibration error (Ec)

was %Em, from the calculation above. Undocumented drift error (Eu) was from the final

correction applied. Because there was only one discrete and an undocumented drift

correction was applied to meet the discrete, discrete error (Ed) was zero for the period.

The grade is “Good” for specific conductance, when Et = 5.24% (Table 7). Additionally,

Et can be entered into the spreadsheet (“Total Error (%)” box, Figure 25) to generate a

schedule of prorated grades for the period. For simplicity, these prorated grades are

typically not applied. Instead, the lowest grade is applied for the entire period. One

exception is when Et results in unusable data. In this circumstance, the division between

poor and unusable is used to isolate only the data that are unusable.

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Final Approval

Final approval is the last step in the data management process that confirms the data

by validating the corrections, grading, and decision making processes of the individual

originally responsible for the data. Final approval is typically conducted by individuals

with the most experience with continuous instream monitoring. In Aquarius, approvals

are applied using the Set Approval action in the data corrections tool box. Division of

Water Quality uses two types of approvals, “Approved” and “In Review”. If a reviewer

disagrees with, or has a question about a correction or grade, the section is marked “In

Review” and the section is discussed with a third party. The third party helps come to a

conclusion on the data management, at which time the data are then approved. Once

data are approved (Figure 27), it may be placed into the report and used in analyses.

Figure 27. Specific conductance data at Aughwick Creek from 2014 displaying the

grades and approval status within the Aquarius data correction tool box. The upper and

lower rows of colored bars at the bottom represent grades and approvals, respectively.

Because the last period was graded unusable (purple), the data were deleted. All

sections were approved (green) except the last section. Since the final reviewer did not

agree with the correction and/or grade, that period of data was marked “In Review”

(yellow) until the discrepancy was resolved with a third party.

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REPORTING

DEP’s Water Quality Division compiles, interprets, and reports instream monitor data in

Continuous Instream Monitoring Reports (CIMR). Continuous instream monitors

produce large amounts of data that, without the proper software, can be difficult to

manage, interpret, and report. Division of Water Quality staff uses both numeric and

visual procedures to report on time series data. Reporting conventions for time series

data often include the maximum, minimum, and median or mean for a measured value

as well as graphical representation of the entire dataset. Data are made available to the

public online after final review and approval are complete. Standard reporting units and

resolutions are detailed in Table 8.

Table 8. Field parameter reporting conventions.

Field Parameter Reporting Units Reporting Resolution

Temperature °C 0.01 °C

Specific Conductance µS/cm 0.1 µS/cm

pH Standard pH units 0.01 standard pH unit

Dissolved Oxygen Concentration mg/L 0.01 mg/L

Dissolved Oxygen Saturation % 0.1 %

Turbidity FNU 0.1 FNU

Aquarius software is used for graphical representation of time series data. When

possible, these graphs will incorporate at least one measured parameter with either a

measured depth or a calculated discharge (Figure 28). Depth or discharge will often

contextualize events of interest (i.e., sudden changes) within the dataset.

A great deal of significance is placed on the relationship of the five water quality

properties and discharge variations, but other event-related changes are equally vital

and can be factored into the relation only through historical measurements, field

experience, and on-site observation. Some examples include but are not limited to

changes in ambient temperature, periods of sustained cloud cover, chemical spills,

increased photosynthetic production, increased wind conditions, sewage overflows,

road construction, and ice formation. Understanding these relations is an essential

element of accurate instream record management (Wagner 2006).

A CIMR not only includes time series data, it also contains any other associated data

collected during the survey period. These data include, but are not limited to, chemical

grab samples, macroinvertebrates, fish, and habitat assessments. When combined,

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these data provide a powerful assessment tool and excellent information on the aquatic

environment.

Figure 28. Depth and pH data for the East Branch Clarion River near the mouth of Gum

Boot Run from September, 2010 to February, 2011. A critical depression in pH can be

seen around the beginning of December, 2010. The depth time series helps identify and

explain that the problem is most likely due to atmospheric deposition through

precipitation.

ARCHIVING RECORDS

Once a report is written and reviewed it will be archived into digital stream files within

DEP and made public online. Monitoring Section personnel within DEP are responsible

for the collection, analysis, manipulation, and storage of instream monitoring data, and

must ensure that the specified requirements of archiving electronic data are fulfilled.

In addition to electronic data, original water-quality monitoring data on paper may

include field notes, field measurements, calibration notes, and service requests. Field

notes and measurements should be included in the appropriate electronic Field Form

for the site that is then archived in the network CIM folder (field measurements are also

stored in Aquarius). Physical calibration logs are scanned and archived in the CIM

folder and all documentation of equipment servicing is logged in the sonde management

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database. It is the responsibility of Monitoring Section to ensure that project files are

organized, complete, and entered into the instream monitor archive. The archive should

be well documented and maintained by appointed personnel in the Monitoring Section

of DEP.

SUMMARY

This protocol offers guidelines for DEP personnel and others in site and monitor

selection considerations, field inspection and calibration procedures, data evaluation

and correction, and data reporting processes. Evolving sensor technology increases the

variety of measurable chemical constituents, allows lower detection limits, and provides

improved stability and accuracy. Protocol guidelines and quality assurance procedures

will continue to be refined as equipment and software progress.

LITERATURE CITED

Anderson, C. W. 2005. Turbidity. Book 9, chapter A6, section 6.7 (version 2.1). in

National field manual for the collection of water quality data: Techniques and

methods. U.S. Department of the Interior, U.S. Geological Survey, Reston, VA.

(Available online at: https://www.usgs.gov/mission-areas/water-

resources/science/national-field-manual-collection-water-quality-data-nfm?qt-

science center objects=0#qt-science center objects.)





Radtke, D. B., J. V. Davis, and F. D. Wilde. 2005. Specific conductance. Book 9,

chapter A6.3 (version 1.2). in National field manual for the collection of water

quality data: Techniques and methods. U.S. Department of the Interior, U.S.

Geological Survey, Reston, VA. (Available online at:

https://pubs.er.usgs.gov/publication/tm9A6.3.)

Ritz, G. F., and J. A. Collins. 2008. Measurement of pH. Book 9, chapter A6.4 (version

2.0). in National field manual for the collection of water quality data: Techniques

and methods. U.S. Department of the Interior, U.S. Geological Survey, Reston,

VA. (Available online at: https://pubs.er.usgs.gov/publication/tm9A6.3.)

Rounds, S. A., F. D. Wilde, and G. F. Ritz. 2013. Dissolved oxygen. Book 9, chapter

A6.2 (version 3.0). in National field manual for the collection of water quality data:

Techniques and methods. U.S. Department of the Interior, U.S. Geological

Survey, Reston, VA. (Available online at:

https://pubs.er.usgs.gov/publication/tm9A6.3.)

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Wagner, R. J., R. W. Boulger, C. J. Oblinger, and B. A. Smith. 2006. Guidelines and

standard procedures for continuous water-quality monitors: Station operation,

record computation, and data reporting. in U.S. Geological Survey Techniques

and Methods. (Available online at: https://pubs.er.usgs.gov/publication/tm1D3.)

Wilde, F. D. (editor). 2004. Processing of water samples. Book 9, chapter A5 (version

2.2). in National field manual for the collection of water quality data: Techniques

and methods. U.S. Department of the Interior, U.S. Geological Survey. (Available

online at: https://www.usgs.gov/mission-areas/water-resources/science/national-

field-manual-collection-water-quality-data-nfm?qt-science center objects=0#qt-

science center objects.)

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4.4 SEDIMENT CHEMISTRY DATA COLLECTION PROTOCOL

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Prepared by:

Amy Williams






2015

Edited by:

Amy Williams






2017

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INTRODUCTION

DEP strives to better understand environmental issues and systems through various

forms of data collection. Water, macroinvertebrates, and fish are routinely collected to

help determine a waterbody’s health and analyze pollution sources and causes.

Streambed sediment is another parameter that can explain a lot about a system.

Bed sediments can accumulate substances that may not be detectable in a single

discrete water sample. They have the potential for accumulating a variety of trace

elements and toxins. Additionally, they could re-enter the water column as suspended

sediment, introducing those substances into the system and causing further problems

downstream. It has been shown that bed and suspended sediments accumulate more

trace metals than are normally present in the water column (Horowitz 1985).

Monitoring streambed sediment parameters can assist in point and non-point source

investigations, spill and complaint issues, and routine monitoring. Testing sediment

could show if contaminants are present, such as high levels of metals, radionuclides, or

organic compounds, and assist in making assessment decisions on impairment or

attainment of flowing waterbodies.

This document provides guidelines for the standardized collection of streambed

sediment samples from flowing waterbody systems. The procedures described here are

adapted from scientific, peer-reviewed procedures, and were developed, field tested,

and implemented by DEP’s technical experts. This protocol does not attempt to

describe the entire spectrum of sediment sample collection techniques, and review of

other documentation is encouraged depending on the specific sampling situation.

Because sampling situations vary largely, no single sediment sampling procedure can

be universally recommended. This document describes sediment sampling procedures

appropriate for typical DEP investigations and may require modification as situations

dictate. Variations to this protocol will be dependent upon site conditions, equipment

limitations, or limitations imposed by the procedure. Investigators should document

modifications and report the final procedures and equipment employed.

Investigators should be aware of, and work to prevent, the potential for sample

contamination at all phases of the sample collection process by observing proper

sample collection, handling, and preservation procedures described here. The most

common sources of error (also known as “interference”) are cross-contamination and

improper sample collection and preservation.

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Field staff required to collect surface water samples must have an assigned four-digit

collector identification number (e.g., 0925). This number along with a sequential three-

digit sample number (e.g., 0925-001), and date/time of sample are used to identify

individual samples. Supervisory staff can request collector identification numbers for

their field staff with the “Collector ID Request Form” found at the eLibrary website (ID

Request Form).

DEP Laboratory Sample Submission Sheet

Field staff will need to have at least Querying and Sample Entry security roles for the

program or business unit that they are collecting samples for. The program or business

unit will be consistent with the Program Code entered on the “DEP Laboratory

Submission Sheet” (Submission Sheet). Collectors must submit samples to the DEP

Bureau of Laboratories (BOL) using a “Sample Submission Sheet”. Field staff are

required to document collector identification number; reason code; cost center; program

code; sequence number; date collected; time collected (military time); fixative(s) used;

standard analysis code (SAC); under certain circumstances, legal seal numbers for

each sample collected; the number of bottles submitted per test suite; collector name;

current date; phone number; and any additional comments that lab analysts will use to

properly handle samples.


member collecting samples. Reason codes, cost centers, and program codes are

program-specific and should be obtained from the program responsible for coordinating

sampling efforts. Sample sequence numbers are three-digit sequential numbers (001-

999) unique to a sample collected on a given day. Date and time collected should be

accurately documented. If a sample is “fixed” or preserved with acid this must be

documented in the appropriate space.

A standard analysis code or SAC is a unique code that details analytical tests to be

applied to a specific sample. Suite codes are also available, and are often used for

radionuclide and organic samples. Each DEP program uses different SACs or suites for

specific projects or purposes. For example, SAC 018 is used by “Water Supply

Management” when submitting water chemistry samples for Special Protection Surveys.

The analytes/tests listed under SAC 018 are those specifically identified by regulation

that surface water must meet and therefore be assessed for if a special protection

determination is warranted. Other programs have developed SACs for their unique

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purposes, and the DEP BOL encourages programs to create SACs tailored to a

program’s specific needs.



analysis. Legal seals are not always needed, and should be used according to a

program’s specific requirements. Legal seal numbers must be singly listed (include

letter and number) for each sample. Legal seals can be obtained from BOL. Refer to

BOL for more information concerning legal seals for the particular needs.

Collector name, current date, phone number, and number of bottles submitted per test

suite were added to the DEP laboratory “Sample Submission Sheet” to meet National

Environmental Laboratory Accreditation Program (NELAP) chain-of-custody

requirements. Using the area at the bottom of the form, each bottle submitted for the

samples identified must be accounted for by enumerating the number of bottles per

category listed for inorganic and organic analyses/tests. Each submitted form is also

required to have printed the collector’s name, current date, collector’s signature

(Relinquished by:), and collector’s phone number. There are also spaces to document a

facility name, facility identification number, and an alternate contact. These three pieces

of information are not required.

The last pieces of information to be documented are any additional comments that lab

analysts need to properly handle samples. This information is documented in the

‘Comment’ field at the bottom of the form. The most common use of this field is to add

or delete tests to or from a specified SAC. For example, SAC 018 does not include a

test for turbidity; however, a sample collector may need to document turbidity for a

particular sample. The sample collector would identify SAC 018 in the appropriate field

and indicate in the ‘Comment’ field to add the turbidity test to the particular sample. If a

large number of samples will be submitted with consistent modifications of a particular

SAC, the BOL prefers a new SAC be created specifically for those samples. Other

important comments to consider include odor, sheen, color, viscosity, foaming, field

meter readings, historical knowledge about the site, and any other information that lab

analysts may need to safely and correctly handle the samples. The BOL requests

collectors contact the appropriate BOL staff before submitting samples requesting

organic tests, potentially dangerous samples, or samples that need to be handled

differently.


DEP programs can and do employ a variety of program-specific sediment sampling

techniques that require a multitude of supplies (sampling bottles, bags, etc.), and can

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include specialized equipment (En Core™ soil corer, Ponar dredge, etc.). This section

describes equipment and supplies required to employ the most commonly used

sampling techniques and does not include all streambed sediment sampling techniques

that could be employed. Additional techniques may be added as they become

applicable and as standard procedures are solidified. Much of the equipment used to

sample soils can be used to sample sediments as long as sediment is not lost in the

water column on the way to the surface; sampling equipment will be determined by

water depth, velocity, and other physical parameters.

Parameter-specific equipment requirements, sample collection, labeling, and storage

will be covered in subsequent sections below. Recommended sampling equipment and

other gear are listed in the below check list:

SEDIMENT SAMPLING

Equipment:

spatulas/trowels/scoops (plastic = metals collecting; stainless steel = organics

collecting)

dredges (e.g., Eckman, Ponar)

corers with vials (e.g., En Core™)

Sample containers:

500 ml plastic sample bottles – trace elements

500 ml amber glass sample bottles – organics

40 ml amber glass vials – organics moisture

determination

plastic bags, 4 Mil thickness - radiological samples

Sample processing:

compositing bowl(s) (plastic = metals collecting; stainless steel = organics collecting)

sieve(s) - < 63 micron

cloth mesh (disposable sieve)

Other

brushes

Munsell color chart

Equipment cleaning:

wash bottle/deionized water

detergent (0.2% phosphate-free)

methanol (to rinse organics equipment)

hexane (to rinse organics equipment)

5% HCl (to rinse trace elements equipment)

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FORMS

sediment field forms

laboratory sediment chem. sheets

SHIPPING

courier shipping forms

tape & dispenser

ice

coolers

Misc.

hip boots

waders

nitrile gloves

markers (black Sharpies), pens, & pencils

GPS/maps

Ziploc® bags (for bottles and/or sample submission sheets)

field meter (e.g., YSI), calibration solutions, spare batteries

calculator

insect repellent

screwdriver/tools

batteries (D-cell, other:________________)

other: _______________________________

The numbers and types of sample bottles field staff need for one sediment sample

depend on the specific SAC or suite. Sample bottle and preservative requirements can

be obtained by contacting DEP’s BOL directly.

The choice to use scoops/spatulas, corers, or dredges depends heavily on-site

conditions and the specific project requirements. Hand-held scoops and spatulas are

convenient for shallow, slow-moving, wadeable streams. They are easy to use and

clean. Unfortunately, with this procedure, it can be easy to lose fine sediment, which is

desirable for most tests. Additionally, it can be more difficult to gather in deeper water,

and if contamination is a concern, it increases the likelihood of skin contact with

material. Deeper and faster-moving water may require the use of a hand-held or

mechanical dredge, such as an Eckman or Ponar. The use of a boat may be required

or, alternatively, dredges can often be lowered from bridges. However, the project aim

should be considered in choosing where to lower a dredge from; runoff from the bridge

may or may not be desired. These are convenient ways to gather bed sediment when

the bottom cannot be manually reached, but a downside is that it is easy to gather too

much deeper sediment, and there is far less control over where and what is actually

collected. If historical sediment information is desired, corers are an option. These

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range from large, cumbersome varieties to small, handheld corers, such as those used

for volatile organic compounds (e.g., En Core™). In all cases, excessive ornamentation

or wooden parts are discouraged to permit easy and thorough decontamination (USEPA

2012). Collectors are encouraged to seek guidance on the variety of dredges and corers

available from the literature at the end of this document.

Some chemical analyses require laboratory technicians to calibrate specialized

laboratory equipment, prepare specialized reagents, or otherwise perform pre-analytical

preparation before samples can be analyzed. If a collector is going to submit several

samples involving specialized preparation, such as allowing radionuclide samples to

ingrow, he or she should contact the appropriate technician at the laboratory to ensure

enough time is allocated for the pre-test procedures. Additionally, large quantities of

samples should always be pre-authorized with the laboratory to avoid surpassing

holding times.

SAMPLING DESIGN CONSIDERATIONS SPECIFIC TO SEDIMENT

Determining sample design is one of the most important parts of a field collection plan.

It is necessary to determine whether a statistical or judgment-based approach is

desired. This depends substantially on the project questions – for example, whether or

not background samples are being collected, or a spill or discharge effect is being

investigated. See Chapter 2, Sampling Design and Planning (Lookenbill 2020) for

suggested sampling plans.

During or after completing a sample collection plan, the collector should perform field

reconnaissance of the site(s). Ease of site access, stream flow, and sediment

deposition locations should be reviewed. One should ideally avoid sampling sediment

immediately after a significant precipitation event where runoff from the surrounding

land could occur (Shelton and Capel 1994). Manmade structures, such as bridges,

should be noted and sampling immediately downstream of them should be avoided.

Runoff from structures can easily impact surrounding sediment downstream. Point and

non-point sources of pollution should be noted. Field reconnaissance is also an

opportunity to determine necessary equipment for sediment collection, such as dredges

versus scoops. Additionally, it can assist in determining locations where it is most likely

for sediment to deposit and where concentrations of contaminants could be highest.

Locations where sediment can accumulate, and therefore accumulate contaminants,

include areas with aquatic vegetation, where velocity decreases, and inside bends in

the stream (Oak Ridge Associated Universities (ORAU) Training 2012). Sediment can

also accumulate near man-made structures, but again, care must be taken to ensure

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sediment in those locations is of interest to the project and not a result of surrounding

land runoff, if that is not the target.



Collectors need to first ensure they have formed an adequate sampling plan that will be

representative of the system under investigation. Care must be utilized during collection

to reduce contamination from outside sources and maximize the integrity of the sample.

The most common causes of sample interference during collection include poor sample-

handling and preservation techniques, input from atmospheric sources, and

contaminated equipment or reagents. Each sampling site needs to be selected and

sampled in a manner that minimizes bias caused by the collection process and that best

represents the intended environmental conditions at the time of sampling.

As stated above, the lab should be contacted prior to large or unique sampling events.

Previous weather conditions and discharge may be analyzed to ensure samples are

being collected with the stream characteristics required to answer the problem in

question. Large precipitation events resulting in high discharge and runoff are not the

best time to sample to judge maximum sediment contamination from a point source

discharge. In addition, rapid water velocities and increased turbidity make sediment

collection with traditional trowels difficult. Discharge data for continuous monitors

throughout the state can be found on the United States Geological Survey website.

Equipment should be gathered and a checklist made prior to departure. Once at the

site, before handling sample containers, the collector should ensure his or her hands

are clean and not contaminated from sources such as food, coins, fuels, mud, insect

repellent, sunscreen, sweat, or nicotine. Gloves should be worn for all sediment

sampling due to the high possibility of sample contamination. Unlike water samples,

sediment samples are handled quite a bit more and there are many more opportunities

to introduce contaminants.

For most sediment sampling, the surficial 1 to 6 cm of sediment with grain sizes < 0.06

mm (silts and clays) are desired. It is best to avoid sands (0.06 – 2 mm) and larger grain

sizes. Fine grained particles tend to accumulate more trace elements than coarser

particles. This is due to a number of factors, including higher available surface area on

finer particles (Horowitz 1985). For some organics analyses, grain size < 2.0 mm (sand,

silt, clay) is acceptable (Shelton and Capel 1994). In general, the way to distinguish

between sand, silt, and clay is displayed in Table 1.

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Table 1. Sediment grain sizes. Adapted from: Ohio EPA. 2001. Sediment sampling

guide and methodologies. 2nd edition.

Soil Type Grain Size Description

Sand 0.06 – 2.0 mm gritty, non-plastic, loose particulates

Silt

0.004 – 0.06

mm

smooth, talc-like, non-plastic, loose

particulates

Clay < 0.004 mm dense, moldable like putty, cohesive

In the event that a site contains large amounts of sandy material, sieving may be

necessary if the aim is to determine the highest concentrations present. In general, it is

a good idea to make sure the sample contains > 30% fine material, i.e., silts and clays

(Ohio Environmental Protection Agency 2001).

All equipment is cleaned prior to being used. It is advisable to wash and soak it for 30

minutes in a 0.2% phosphate-free detergent (Shelton and Capel 1994). After soaking,

rinse the equipment thoroughly in deionized water, air dry (if possible), and store in

individual, sealable containers or bags. If sampling for organic contaminants, clean the

stainless steel or TeflonTM equipment with soap and tap water, rinse with deionized

water, dry, rinse with acetone, and, lastly, rinse with hexane (as recommended by DEP

BOL staff). Wrap organics equipment in aluminum foil. It is highly advised to have

several sets of equipment, particularly if planning to sample reference (background) and

impacted sites in one trip. It is difficult to fully clean equipment in the field. If doing so,

save the rinsate and dispose of it properly. It is also advisable to occasionally collect a

rinsate blank.

First, calibrate any field sampling equipment, such as field meters according to Chapter

4.1, In-Situ Field Meter and Transect Data Collection Protocol (Hoger 2020). If dealing

with a possibly impacted site and reference location(s) have been chosen, sample

reference locations first if combining impacted and reference sites into one trip. This will

help lessen the possibility of contamination between sites. It is important to note that

safety is always first – avoid wading into high flow areas or locations where slippage

could occur, wear appropriate wading gear, and have the proper length and type of

gloves on if dealing with impacted sites or chemicals.

If sampling is taking place due to a point-source pollution incident, it is advised to

sample downstream of the discharge, at the discharge, and upstream of the discharge,

at minimum. In order to collect a sample representative of a particular location, plan to

collect at between 5 to 10 depositional zones within a study site and composite those

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into one sample (Shelton and Capel 1994). For example, if collecting at a point of acid

mine discharge (AMD), composite sediment at between 5 to 10 depositional zones

downstream of the discharge, composite sediment at between 5 to 10 depositional

zones at the discharge, and composite sediment at between 5 to 10 depositional zones

upstream of the discharge. Compositing several “zones” of sediment will allow for a

more representative description of the sample area and will not target one spot.

Additionally, due to lack of sediment buildup at some locations, compositing several

spots of sediment buildup facilitates obtaining only the first 1 to 6 cm of sediment. Of

course, specific project needs may deviate from this, so plan and document the study

accordingly. Additionally, composite sampling is not appropriate for the collection of

volatile organics samples because compounds could be lost.

Sampling stations located upstream of the discharge pipe should be in non-impacted

areas to serve as controls. If there are multiple discharges, then sample stations should

be placed to bracket individual discharges in order to better characterize each source.

For sampling downstream of the discharge pipe, if the investigator is interested in

determining the downstream point where the discharge ceases to be an effect in the

sediment, the investigator should avoid the immediate vicinity of the discharge/influence

point and select a sample point far enough downstream to allow for mixing between the

discharge and stream flow. Sampling can occur at any noted deposition points between

in order to characterize effects of the discharge. Depending on stream size, flow,

velocity of the plume, and angle at which it enters the stream, solids may deposit at a

variety of locations.

As a summary, composite-sampling 5 to 10 depositional zones in a single location

(upstream of a discharge, for example) is recommended in most cases. In addition, the

sampling reach is discretionary and based upon site-specific characteristics and

questions being answered. Regardless of the technique implemented, similar sampling

procedures should be used if sampling several locations in one single study, to allow for

comparability of data.

At each site, always collect water samples first to avoid stirring up sediment that could

contaminate the water sample. Take any water field measurements before sediment

sampling. Additionally, always work in a downstream to upstream fashion at a site.

Lastly, always document the appearance, odor, sheen, and feel of the sediment on the

‘Sediment Field Data Collection Sheet’ (see Appendix B.4).

Wadeable Samples

Gather the equipment (scoop, appropriate sampling container [see parameter-specific

instructions, below], gloves) and enter stream at the most downstream of the sampling

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locations. If interested in surficial, recent deposition, sample only the top 1 – 6 cm of

sediment; if interested in historical sediment accumulation, sample > 6 cm. Facing

upstream, rinse the sampling container several times with native water, emptying it

behind you (downstream) to avoid stirring up any additional sediment. In addition, rinse

all sampling equipment in native water as well, including scoops, sieves, and bowls.

Enter water column at the first sediment depositional area, using care to gather

upstream of the current standing position and not stepping where collection will occur,

and gather a scoopful of sediment. Slowly raise the scoop out of the water to avoid

losing fine material and place sediment into sampling container. Move to the next

depositional zone and repeat, working from downstream to upstream. If too much water

is collected, carefully decant as it is collected, trying to allow it to settle first as much as

possible to avoid loss of fine materials. Additionally, avoid collecting vegetation or other

debris. The lab needs to pulverize samples before analysis, so the finer sediment and

the less debris collected, the better.

Once enough sediment is collected, exit the stream and empty the sample into an

appropriate compositing container. Remove any large pebbles, sticks, vegetation, or

other debris. Allow the sediment to settle and carefully decant the water off, or,

alternatively, sieve the sample to ensure only fine grained materials are collected.

Generally, if one sample is sieved in a project, all samples in that project should be

sieved for consistency. In any case, fully note on the field sheet the texture, color, and

odor of the sediment. Homogenize the sample using a spatula or trowel. One

recommendation to fully homogenize is to “quarter” the sample into four parts, mix those

parts, and then mix the entire sample together (USEPA 2012). Place sample back into

the sampling container and record all details about the site and collection procedures on

the field sheet. Add fixative to the sample, if required. If refrigeration is necessary, place

the sample in a cooler with ice in order to cool the sample to 4°C. Do not use dry ice

because the sample may freeze. Record GPS coordinates of the location and take

photographs, if necessary.

Corers may be necessary if the interest is in collecting historical sediment or volatile

organic compounds that cannot be exposed to air. Coring allows stratification of the

sediment and the possibility of testing individual layers. A corer is pushed into the

sediment and the core is captured in the sample tube (ORAU Training 2012). They can

be particularly useful in fast-moving streams where grab samplers would be difficult to

use. They come in a variety of shapes and sizes for different needs. Many include a

valve or catcher that will prevent loss of the core as it is being brought to the surface.

The DEP BOL requires the use of an En Core™ corer (or similar) or a corer using pre-

preserved (methanol) vials for volatile organic compound soil and sediment samples. If

it is not possible to use these types of corers, a note must be made on the “Sample

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Submission Sheet” or a data release form will be sent. USEPA has a detailed

description on how to collect volatile organic compounds in USEPA Test Method 5035A

(2002) - Appendix A. Refer to the DEP BOL for more information regarding volatile

organic compound collecting.

To collect sediment with a corer, either push or hammer it into the sediment (ORAU

Training 2012). Be sure to push it straight down and pull it straight up without turning it,

and do not fill the tube completely. Keep the sampler vertical when pulled out of the

sediment to avoid mixing. Cap the bottom of the sample tube as soon as possible, then

the top. If there is some water left in the tube, carefully decant it off and stuff a cloth into

the end – this reduces mixing. Augers, often used for soil, can be used for collecting

sediment and are turned and pushed into sediment; however, sample loss could be an

issue. Refer to the references in the “Literature Cited” section of this document for

further information on using a variety of corers, augers, and dredges to collect

streambed sediment.

Nonwadeable Samples

If the water flow is too deep and/or fast moving to manually collect sediment, consider

using a dredge. These can either be lowered from a boat or a structure such as a

bridge. Depending on size, they can be manually lowered or lowered using a winch. If

boating, turn off the engine before using the dredge to eliminate impact on the sample.

If an anchor is used, make sure dredging is not collecting the sediment that the anchor

disturbed.

Once at the desired location, set the dredge according to user instructions and carefully

lower it to the stream bottom. Trip the dredge, and slowly bring it to the surface. Avoid

lowering and raising it too fast, which risks disturbing the sediment before it reaches the

bottom or losing sediment on the way up. Once at the surface, examine the sample and

determine if the dredge was properly tripped; if it was not, discard the sediment and re-

sample. Dispense the sample in an appropriate compositing container, collect the rest

of the depositional zones, and follow the instructions above for decanting/sieving and

compositing the sample. See the references in the “Literature Cited” section of this

document for further information on using a variety of dredges to collect streambed

sediment.

Parameter-Specific Considerations: Radionuclides

Stainless steel, high-density polypropylene (HDPP), or polyethylene (HDPE) sampling

equipment is recommended for sampling for radionuclides (USEPA 2012). Sampling

containers should also be made of the same plastics and have a polytetrafluoroethylene

(PTFE) or TeflonTM lined lid (USEPA 2012). Verify that the lid will not absorb any water.

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Stainless steel or plastic compositing containers are recommended. If testing for

gamma-emitting radioisotopes, bags are recommended due to large sample volume.

DEP BOL recommends collecting one (1) kg of wet sediment in order to verify having

enough sample to analyze ½ kg of dry sediment in the lab. Two 500-mL bottles are also

a sufficient amount of sample for analysis. Double or triple-bagging samples is highly

recommended. Properly close the bags and use a non-absorbing tape to seal. Gamma-

emitting radioisotope sediment samples do not require refrigeration or preservatives.

The sample holding time is 72 hours if iodine analysis is needed, otherwise it is 6

months. For other tests and details, contact the DEP BOL.

Parameter-Specific Considerations: Metals

High-density polypropylene (HDPP) or polyethylene (HDPE) sampling equipment is

recommended; avoid metal sampling equipment whenever possible. There is always

the chance of metals in the equipment influencing sample results. Plastic compositing

containers are recommended. Most standard plastic water-sampling bottles can be

used to store sediment for metals testing. A typical sediment metals sample at DEP

BOL requires a 500-mL plastic or glass container. No preservatives are required and

refrigeration to 4°C is necessary. Holding time is 6 months. Contact the DEP BOL for

specific parameter details.

Parameter-Specific Considerations: Volatile & Semi-Volatile Organic Compounds

As stated above, DEP BOL requires samples for volatile organics to be collected with

corers. There are two coring procedures available: The first is the En Core™ - for each

analysis, 2 x 5g En Core™ vials are needed, plus one (1) 40mL amber glass vial, to be

used for moisture determination. There is also a 25g size available, but this is not

recommended and will also require a data release form. A second procedure uses a

small, disposable corer that contains pre-preserved (5mL of methanol), pre-weighed

vials. Weight is recorded on the outside of the vial. The core is collected, the sample is

ejected into the vial, and the vial is quickly capped. The vial is not to be packed full and

care should be used to avoid losing methanol. The threads of the vial should be cleaned

before capping so methanol does not leak out. Do not attach additional labels to the vial

since this will change the weight of the vial. Two pre-preserved vials and one (1) 40mL

amber glass vial are required for analysis. Only “high concentration” vials are required.

If unable to use these types of samplers, specifically state the reasons why on the

“Sample Submission Sheet” or a “Client Request for Data Release” form will be sent. In

the instance of not using a core sampler, volatile organics samples need to be packed

tightly in 2 x 40mL amber bottles (unpreserved). A recommended procedure to avoid air

contact is to take a scoop of sediment, remove the top layer in the scoop, and

immediately fill the vial with sediment from the scoop. Samples have a holding time of

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48 hours using the En Core™, and must be received by the lab within 24 hours after

collection. Samples collected and preserved with methanol have a holding time of 14

days. See USEPA Test Method 5035A (2002) - Appendix A for more details.

If testing for semi-volatile organic compounds, fill a 500mL amber glass sampling jar.

Volatile and semi-volatile organics sediment samples need to be refrigerated to 4°C and

do not require field preservation, unless collecting volatile organics in the pre-preserved

vials. When possible, always schedule with the lab before collecting samples for organic

analyses.

Labeling

While the minimum requirement is the collector number and sequential sample number,

collectors are encouraged to add date and time collected, general test(s) description

(total metals, etc.), and preservation indication. This will help prevent confusing what

bottles are from which tests and to help ensure the sample is properly preserved and

stored. Labeling should be done so that at least 1” of space is left at the top of the bottle

to allow BOL to apply lab labels.

Permanent marker will rub off HDPE bottles during collection and transport. Therefore,

clear packing tape should be wrapped around the bottle to protect hand-written labels.

BOL discourages the use of masking tape. Collectors should keep a log book of all

samples they take, and should not re-use sequential numbers in order to avoid

confusion. The sample log should annotate the unique collector identification and

sample number, date and time, the waterbody name, sample location, SAC code, and

any additional analytical tests performed or excluded.

Quality Assurance

A duplicate grab sample should be collected every 20 samples or for each sampling

trip/day to gauge testing variability and potential sources of contamination due to

collection procedures. Duplicate samples are collected simultaneously or sequentially

with the associated environmental sample, using identical sampling and preservation

procedures. Sequentially collected duplicates may measure inhomogeneities present in

the sediment. Duplicates are assigned unique sequential sample numbers. The

collector needs to carefully annotate which sample is a duplicate in their field book.

Duplicates must be documented appropriately in SIS under the ‘Comments/Quality

Assurance’ tab.

“Split samples” are collected as one sample and then divided in two for separate

analysis (Radtke 2005). These can be sent to different labs for analysis or be tested at

the same lab, and can help determine lab analysis variability.

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“Rinsate blanks” may be necessary at times if a highly-impacted site was sampled. After

cleaning equipment, a sample of the cleaning rinsate water is saved and analyzed for

parameters of concern (USEPA 2012). This verifies whether or not the equipment was

properly cleaned.

DEP BOL does not require sediment or soil blanks, although they can be submitted.

Normally these are purified sand or water.

Post-Sampling Decontamination

If sampling multiple locations in one trip, try to have multiple sets of equipment, since

decontamination in the field can be difficult. If only one set of equipment is available and

multiple locations must be sampled, decontamination must occur in the field. Whether

completed in the field or back at the office, wash all used equipment first in phosphate-

free detergent. Brushes are helpful for removing caked-on soil. Rinse thoroughly. Then,

rinse all non-plastic equipment with methanol. Rinse non-metallic equipment, such as

those used to sample metals, with a dilute acid, such as 5% HCl (Radtke 2005, Shelton

and Capel 1994). It may be necessary to rinse equipment used to sample for

radioisotopes with a radioactive decontaminant, such as NoCount®. Afterwards, rinse

all equipment thoroughly with deionized water and allow to air dry (if possible).

Equipment that is used to sample for organic contaminants should be rinsed with

acetone and then hexane, rather than methanol or acid, after the soap and water wash.

Discard disposable equipment rather than clean it. Always be sure to store

contaminated/dirty equipment separately from unused/clean equipment. Store

equipment in individual, sealable containers or plastic bags. Store equipment used to

sample organics in aluminum foil.


Samples need to be shipped or delivered to the lab as soon as possible. The collector

should understand that certain laboratory analyses have “holding times” during which

tests must be conducted for result validity. Volatile organics, for example, must be

received by the laboratory within 24 hours after collection. If a sample surpasses

holding time requirements the results will not be reported unless a “Client Request for

Data Release” form (see the BOL website) is submitted to the Bureau of Laboratories. It

is not advisable to collect and ship samples on Fridays, as the laboratory does not

operate on weekends; samples shipped on Friday will not be received and logged until

Monday morning. Collectors essentially need to plan their sampling from Monday

through Thursday and verify the samples will reach DEP BOL by early Friday morning

at the latest. The days before holidays will also need to be considered.

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Shipping

All DEP district and regional offices are designated pick-up locations for water and

sediment samples. In most cases, samples must be dropped off for pick up by 1600

hours. Other locations exist, such as at some Pennsylvania Department of

Transportation facilities and private businesses, but these drop-off locations may require

call-ahead notice to the current courier, as they may not be visited daily. Further, the

drop-off locations may require a drop-off specific key to open the drop-off entrance lock.

Collectors need to coordinate with the current courier for specific drop-off and pick-up

requirements.



to the cooler to ensure samples remain at 4°C during overnight shipping. Laboratory


samples may be subject to a data release request. Dry ice will freeze samples and

should never be used for storage or shipping. The “Sample Submission Sheet” should

be filled out, inserted into a Ziploc® bag, and attached to the inside of the cooler lid.

Courier shipping labels should be printed out during ordering so they can be attached to

the top of the cooler lid during sample drop-off. Shipping labels are secured to the

cooler lip with two pieces of packing tape on the left and right side; taping all sides of

the label makes removal difficult for lab technicians. Be sure that any required legal

seals are in place.

LITERATURE CITED





Horowitz, A. J. 1985. A primer on trace metal-sediment chemistry. Water-Supply Paper

2277. U.S. Geological Survey, Alexandria, Virginia.





Oak Ridge Associated Universities (ORAU) Training. 2012. Environmental Monitoring

for Radiation. Oak Ridge, Tennessee.

Ohio Environmental Protection Agency. 2001. Sediment sampling guide and

methodologies. 2nd edition. Division of Surface Water, Columbus, Ohio (Available

from: http://www.epa.ohio.gov/portals/35/guidance/sedman2001.pdf).

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Radtke, D.B. 2005. National field manual for the collection of water-quality data,

Chapter A8: Bottom-material samples. Techniques of Water-Resources

Investigations. Book 9: Handbooks for Water-Resources Investigations. U.S.

Geological Survey, Reston, Virginia.

Shelton, L.R. & P.D. Capel. 1994. Guidelines for collecting and processing samples of

stream bed sediment for analysis of trace elements and organic contaminants for

the national water-quality assessment program. Open-File Report 94–458. U.S.

Geological Survey, Sacramento, California.

USEPA. 2002. Method 5035A – Closed-System-Purge-and-Trap and Extraction for

Volatile Organics in Soil and Waste Samples. Appendix A: The collection and

preservation of aqueous and solid samples for volatile organic compound (VOC)

analysis.

USEPA. 2012. Sample collection procedures for radiochemical analytes in

environmental matrices. U.S. Environmental Protection Agency, Office of

Research and Development, National Homeland Security Research Center,

Cincinnati, Ohio.

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4.5 PASSIVE WATER CHEMISTRY DATA COLLECTION PROTOCOL

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Prepared by:

Amy Williams






2013

Edited by:

Amy Williams






2017

Disclaimer:

Mention of trade names or commercial products does not constitute endorsement or

recommendation for use.

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INTRODUCTION

The Pennsylvania Department of Environmental Protection (DEP) often utilizes unique

and innovative techniques to explore environmental issues and determine if

environmental problems exist. One technique that has been employed in recent years is

the use of passive sampling devices to collect water quality samples of low-level

environmental contaminants.

Some contaminants, such as hormones and other endocrine-disrupting compounds

(EDCs), can have detrimental effects on aquatic life at extremely low levels (in the ng/L

range). In addition, it is very easy to miss a time period where a low-level compound

may be detected, which a passive sampler would capture. Therefore, using a sampling

device to collect compounds in a large volume of water over a period of time (days) is a

great way to collect low-level or sporadically-present compounds.

Although this document refers to the use of passive samplers in surface water, they are

also used to sample groundwater and air. There are many types of passive water

samplers available today. The samplers employed by the Water Quality Division have

been the Semi-Permeable Membrane Device (SPMD) and the Polar Organic Chemical

Integrative Sampler (POCIS). SPMDs are used for sampling neutral organic compounds

that have a log octanol-water partition coefficient (Kow) that is greater than 3 (Alvarez

2010). Examples of compounds that can be sampled with SPMDs are organochlorine

pesticides, polychlorinated biphenyls (PCBs), polybrominated diphenyl ethers (PBDEs),

and polycyclic aromatic hydrocarbons (PAHs). A POCIS is used to sample water

soluble organic compounds with a log Kow less than 3. Examples of compounds that can

be sampled with POCIS are various wastewater indicators, many currently-used

pesticides, pharmaceuticals, and hormones.

This document provides guidelines for the standardized collection of passive water

samples from flowing waterbody systems. The procedures described here are adapted

from scientific, peer-reviewed procedures, and were developed, field tested, and

implemented by DEP’s technical experts. This protocol does not attempt to describe the

entire spectrum of passive sample collection techniques, and review of other

documentation is encouraged depending upon the specific sampling situation.

Because sampling situations vary largely, no single passive sampling procedure can be

universally recommended. This document describes passive sampling procedures

appropriate for typical DEP investigations and may require modifications as situations

dictate. Variations to this protocol will be dependent upon site conditions, equipment

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limitations, or limitations imposed by the procedure. Investigators should document

modifications and report the final procedures and equipment employed.

Investigators should be aware of, and work to prevent, the potential for sample

contamination at all phases of the sample collection process by observing proper

sample collection, handling, and preservation procedures described here. The most

common sources of error (also known as “interference”) are cross-contamination and

improper sample collection and preservation.


Sampling Supplies

This section describes equipment and supplies required to employ POCIS and SPMD

sampling techniques and does not include all passive water sampling techniques that

could be employed. Additional techniques and equipment may be added as they

become applicable and as standard procedures are solidified or altered. Very little

equipment is needed to deploy a passive sampler; the sampler, a canister or holder for

it, gloves, and a securing pin are all that is necessary. The following supplies are

recommended:

• Passive sampler(s), already assembled and in a tin until deployment in the field

• Canister or other holder for passive sampler(s)

• Nitrile Gloves (material cannot be composed of any compounds to be tested)

• Stake/pin for securing in streambed

• Carabineers and eye bolts for securing passive sampler holder to stake/pin

• Sledge hammer

Samplers should be pre-assembled and stored in a tin or other container in a

refrigerator (POCIS) or freezer (SPMD or POCIS). New paint can tins are perfect

containers because they seal tightly; no air should be allowed to escape or get in.

Sampler contamination pre-deployment could be an issue; therefore, assemblage in a

sterile environment is necessary.

Factors such as temperature and biofilm buildup can affect the samplers’ ability to

collect chemicals. Performance reference compounds (PRCs) can help alleviate this

problem (Alvarez 2010). These are compounds added to the samplers pre-deployment,

and are measured after deployment to see how much chemical is lost. Adjustments can

then be made to targeted chemical measurements based on the amount of loss

measured in the PRCs. Photo-sensitive PRCs can also be used if the compounds of

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interest are prone to photo-degradation and the samplers will be exposed to direct

sunlight.

Field blanks are also required. Blanks are just regular samplers that will be exposed to

air in the field while the passive samplers are being deployed and retrieved from the

water. Any air contamination will be picked up and measured by these blanks. Air

contamination is common, particularly with SPMDs, depending on the compounds

measured. Industrial areas and cities may produce airborne contaminants that are

easily picked up by SPMDs. The number of blanks implemented is up to the collector’s

discretion – DEP normally uses one blank per three sites/deployments, but more blanks

are always optimal. The only issue with this procedure is that any contamination that

results will be unable to be pinpointed to any one site or sampler.

Analysis Lab(s)

Since many of the compounds that may be analyzed by passive water samplers are

emerging contaminants, there are a limited number of labs that may be able to analyze

extracts. DEP BOL may be able to perform some analyses. Past analyses that have

been performed by BOL on passive sampler extracts include PAHs, pesticides, and

PCBs. DEP will also use USGS and private laboratories to analyze additional

parameters.

SITE SELECTION AND SAMPLING DESIGN

A sample collection plan is recommended before commencing a sampling project. This

should include the location(s) to be sampled, reasons for sampling, number of samples,

media of interest (water, sediment, soil, macroinvertebrates, etc.), parameters to

sample, QA/QC plans, equipment, preservatives, sample container types, estimated

costs, maps, and any other notes or comments pertaining to the project. More

information on developing Project Plans can be found in USEPA 2002a, USEPA 2002b,

and USEPA 2006. It can be very helpful and more descriptive of a site to take grab

water samples, at a minimum, in conjunction with sediment sampling. Macroinvertebrate

and fish samples can also help describe overall impacts to a system regardless of the

chemicals of interest or the problem being investigated.

Prior to sampling, any historical information, aerial photography, and/or topographic

maps of the locations should be gathered and analyzed. Geomorphology and

underlying geology around the waterbody may be helpful in determining parameters of

interest. Point and non-point source pollution possibilities should be documented. Past

water quality and/or biological monitoring results should be gathered. Additionally, it

should be noted whether sampling locations are in close proximity to gaging stations or

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other continuous monitoring stations – this data could prove useful during final data

analysis.

Determining sample design is one of the most important parts of a field collection plan.

It is necessary to determine whether a statistical or judgment-based approach is

desired. This depends substantially on the project questions – for example, whether or

not background samples are being collected, or a spill or discharge effect is being

investigated. Many sampling design options exist, with many options being statistical in

nature. These include systemic sampling, where the distance between sampling

locations is kept consistent (Horowitz 1985). This is not appropriate if a known

contaminant is present in an area of sediment. Another statistical approach is random

sampling, where sample locations are arbitrarily selected so that one area is not more

likely to be chosen over another. If the area is homogenous, this is recommended

(Horowitz 1985). Stratified random sampling is another option. This involves dividing a

target area that may have a contaminant or area of interest into several areas that are

homogenous. Random sample sites are then chosen within those areas. This helps

remove heterogeneity at a site (Horowitz 1985). A non-statistical design approach is

judgmental or targeted sampling – sampling sites are chosen based on professional

judgment, not scientific theory. This is a good approach to use for sites where a known

contaminant is present (i.e., discharge pipe) and can be extremely resource-efficient if

there is a lack of funds or equipment.

It can be helpful to anticipate how the data will be used. Consider if it will impact legal or

regulatory actions, and how the data will be analyzed and presented. Also consider

what criteria may be used during its interpretation. DEP does not have narrative or

numeric passive sampler criteria at this time, but does have numeric water quality

criteria for many chemicals that may be sampled. One can refer to 25 Pa. Code Chapter

93 for details. One thing to keep in mind is that these chemicals are being sampled over

a long period of time, normally at least a month, so the criteria should not be applied as

written since they are normally based on grab samples.

Some SACs/suites are available for passive sampler extract analysis from DEP BOL.

Contact DEP BOL organic section for more details and to set up a sample submission

schedule.

During or after completing a sample collection plan, the collector should perform field

reconnaissance of the site(s). It’s a good idea to view the site or deploy at a lower flow,

particularly if rain or increased flow is expected in the weeks after deployment. If

samplers are deployed at high flow, low flow levels should be approximately known.

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Samplers cannot be exposed to the air during low flow conditions or sample results will

be compromised.



Collectors need to first ensure they have formed an adequate sampling plan that will be

representative of the system under investigation. Care must be utilized during collection

to reduce contamination from outside sources and maximize the integrity of the sample.

The most common causes of sample interference during passive sampler collection

include input from atmospheric sources and contaminated equipment. Each sampling

site needs to be selected and sampled in a manner that minimizes bias caused by the

collection process and that best represents the intended environmental conditions at the

time of sampling.

Samplers should be stored in their containers under refrigerated or frozen conditions

until deployment. SPMDs must be kept frozen (to -20°C, or the temperature that the

laboratory requires) until deployment. POCIS can be stored at ambient temperature for

up to two weeks until deployment; any longer than that and they should be frozen (to -

20°C, or the temperature that the laboratory requires). Once in the field, nitrile gloves

should be put on before sample containers are opened. Blank samplers should be

opened and exposed to air. Samplers should be placed in canisters/holders in a timely

fashion and moved to the water where stakes have been pre-set. Samplers should be

fastened to stakes using carabineers, with careful attention paid to how noticeable the

canisters are. Canisters can be supported with rocks but care should be used to avoid

covering them entirely so that water flow is still possible. Samplers should not be placed

directly in areas of large sediment deposits, if possible, or sediment will clog the

filters/membranes. If the samplers are too visible from shore or from bridges, tampering

of the equipment could occur. Samplers also cannot be exposed to air during periods of

low flow – lowest possible flow during the time period of deployment should be

anticipated. Stakes should be very securely hammered into sediment, or high flows

could loosen samplers. Blanks should be tightly closed after deployment and stored

frozen until sampler retrieval. They should be labeled appropriately so they are used at

the same site(s) during retrieval.

The number of passive samplers needed for any one sample will depend on the

analyses desired (Alvarez 2010). Discuss this with the analysis lab(s) and equipment

lab(s).

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Date and time of deployment, as well as location site conditions (such as pH,

temperature, conductivity, and flow) should be noted. Passive samplers are designed to

be deployed for days to months, although 30 days is the average time (Alvarez 2010).

Short deployments reduce the advantages of using passive samplers; too long of a

deployment could compromise the data. DEP normally deploys passive samplers for

around 30 days, or one month. At the time of retrieval, date, time, and field conditions

should be noted again. The same blanks used during deployment should be air-

exposed and samplers should be removed from the water with the same caution as

deployment. If available, new blanks could be used. The passive samplers should be

placed immediately into containers that remained sealed during deployment (to avoid

contamination). Samplers should then be frozen until received by the lab. Samplers

should be extracted as soon as possible or mailed overnight on ice (packs, not loose

ice, to avoid water contamination as the ice melts) to a lab for extraction. Analysis

should occur in the specified timeframe of holding times of the compounds of interest.

Labeling

Container labeling is not necessary until samplers are retrieved. Be sure to put POCIS

in POCIS tins and SPMDs in SPMD tins since solvents may remain in the tins. Once

they are retrieved, label the containers with sampling locations, dates, and times. Label

blanks accordingly.

Quality Assurance

Because of the expense of deploying passive samplers, DEP does not often do

duplicates as it would other samples; however, at least one duplicate per set of around

10 is recommended. Blanks are described above.

Post-Sampling Decontamination of Equipment

After deployment, passive sampler canisters/holders need to be decontaminated. They

should be scrubbed with non-ionized soap and water to remove sediments and organic

contaminants. If organic compounds were sampled, containers should be rinsed with

acetone to remove any water and then rinsed with hexane (Environmental Sampling

Technologies SOP E-13). Soaking canisters briefly in a solution of hydrochloric acid (2-

4N) with hot water will remove rust and calcium.


Holding times depend on the compounds analyzed. Discuss with the analysis lab(s)

regarding holding times.

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Shipping

Samplers need to be processed as soon as possible after retrieval or shipped as soon

as possible to the lab performing the processing. Samplers will need to be shipped

priority overnight in coolers with ice packs. No loose ice should be used, as it may melt

and get inside the containers, compromising the samples. Processed extracts should be

mailed immediately to the labs performing the analyses.

LITERATURE CITED

Alvarez, D. A. 2010. Guidelines for the use of the semipermeable membrane device

(SPMD) & the polar organic chemical integrative sampler (POCIS) in

environmental monitoring studies. Techniques and Methods 1–D4. U.S. Geologic

Survey, Reston, Virginia.

Environmental Sampling Technologies. SOP E–13, Cleaning of SPMD/POCIS

Deployment Devices. Environmental Sampling Technologies, St. Joseph, MO.

Horowitz, A. J. 1985. A primer on trace metal-sediment chemistry. Water-Supply Paper

2277. U.S. Geological Survey, Alexandria, Virginia.

USEPA. 2002a. Guidance on Choosing a Sampling Design for Environmental Data

Collection for Use in Developing a Quality Assurance Project Plan. EPA QA/G-

5S. U.S. Environmental Protection Agency, Office of Environmental Information,

Washington, DC.

USEPA. 2002b. Guidance for quality assurance project plans. EPA QA/G-5. U.S.

Environmental Protection Agency, Office of Environmental Information,

Washington, DC.

USEPA. 2006. Guidance on systematic planning using the data quality objectives

process. EPA QA/G-4. U.S. Environmental Protection Agency, Office of

Environmental Information, Washington, DC.

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4.6 SAMPLE INFORMATION SYSTEM (SIS) DATA ENTRY PROTOCOL

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Prepared by:

Dustin Shull






2017

4-122

INTRODUCTION

SIS is an Oracle® application that DEP collectors use to store, manage, and retrieve

chemical sample data, except for continuous physicochemical data. Continuous

physicochemical data is stored and managed using a separate system due to the

structure of the data, which is not handled by SIS. This protocol outlines the data

requirements and documentation procedures for handling chemical data after it has

been collected in the field.

SIS REQUIREMENTS

Sample collectors, at the very least, must have security roles for their program to

perform Sample Entry and Querying. Collectors must obtain a SIS login name and

password to enter/edit sample information. Collectors will also need to contact a system

coordinator or eFACTS coordinator to obtain the correct SIS securities (see SIS

Security Request Form, doc # 1300-FM-IT1016 SIS), which will allow them to manage

sample information in SIS. SIS securities are broken down into roles. Roles include (1)

Querying, (2) Project Entry, (3) Monitoring Point Entry, and (4) Sample Entry. Each role

is then applied to one of 52 specific programs or business units such as (1) Watershed

Conservation, (2) Water Supply Management, (3) Land Recycling and Waste

Management. Program or business unit names periodically change, pending

reorganizations and other circumstances.

Samples submitted to the BOL will have the following information populated in SIS:

collector identification number, sample sequence number, sample time and date, and

sample results. It is the responsibility of the collector to populate, at the minimum,

sample medium, sample collection location, field parameters, quality control

identification, and general comment information.

SIS can be accessed through the DEP Intradep website by selecting ‘Oracle

Applications’. DEP maintains several Oracle applications, so users must select ‘SIS -

Samples Information System’. Users will be prompted to enter a unique Commonwealth

of Pennsylvania (CWOPA) username and password, in addition to a database identifier.

The database identifier is ‘prod’. Samples can be entered into SIS by the sample

collector before or after BOL populates sample results. It is important to enter the

collector identification number, sample sequence number, and date and time collected

correctly. If samples are entered into SIS before BOL populates sample results these

attributes will be used to associate sample results. If samples are entered after BOL

populates results, the sample collector will need to query to find the sample and

populate attributes. Additional information and step-by-step instructions are available

through the Sample Information Users Guide (Appendix B.3).

5-2

5.1 STREAM HABITAT DATA COLLECTION PROTOCOL

5-3

Prepared by:

Tony Shaw






2002

Edited by:

Josh Lookenbill






2017

5-4

INTRODUCTION

DEP has adopted a modification of the habitat evaluation procedures outlined in

USEPA’s Rapid Bioassessment Protocols (Barbour et al. 1999). DEP riffle/run stream

habitat is evaluated based on 12 parameters rather than the 10 parameters of Barbour

et al. (1999) and multihabitat stream habitat is evaluated on nine of the 10 parameters,

dropping channel sinuosity of Barbour et al. (1999). The matrix used to assess habitat

quality is based on key physical characteristics of the waterbody and surrounding lands.

All parameters evaluated represent potential limitations to the quality and quantity of

instream habitat available to aquatic biota. These, in turn, affect community structure

and composition. The main purpose of the habitat assessment is to account for the

limitations that are due to existing stream conditions. This is particularly important in

cause/effect and cumulative impact studies where the benthic community at any given

station may already be self-limited by background watershed and habitat conditions or

impacts from current landuses. In order to minimize the effects of habitat variability,

every effort is made to sample similar habitats at all stations. The habitat assessment

process involves rating twelve parameters for riffle/run prevalence waterbodies or nine

for low gradient waterbodies as excellent, good, fair, or poor, by assigning a numeric

value (ranging from 20 - 0), based on the criteria included on the Habitat Assessment

Field Data Sheets (Appendix C) and the information provided below.

The habitat assessment parameters used in the evaluations for riffle/run prevalent

waterbodies as well as for low gradient waterbodies are identified and discussed below.

The first six parameters evaluate stream conditions in the immediate vicinity of the

benthic macroinvertebrate sampling reach.

• Instream Fish Cover (riffle/run & low gradient)

Evaluates the percent makeup of the substrate (boulders, cobble, other rock

material) and submerged objects (logs, undercut banks) that provide refuge for a

variety of fish including both large bodied pelagic species as well as smaller

benthic specialists.

• Epifaunal Substrate

(riffle/run) – Evaluates riffle quality, i.e., areal extent relative to stream width and

dominant substrate materials (cobble, boulders, gravel) that are present.

(low gradient) – Evaluates the relative quantity and variety of natural structures in

the stream, such as large rocks, fallen trees, logs and branches, and undercut

banks.

• Embeddedness (riffle/run)

Evaluates the extent to which rocks (gravel, cobble, and boulders) and snags are

covered or sunken into the silt, sand, or mud of the stream bottom. The rating of

5-5

this parameter may be variable depending on where the observations are taken.

To avoid confusion with sediment deposition (another habitat parameter),

observations of embeddedness should be taken in the upstream and central

portions of riffles and cobble substrate areas.

• Pool Substrate Characterization (low gradient)

Evaluates the type and condition of bottom substrates found in pools. Firmer

sediment types (e.g., gravel, sand) and rooted aquatic plants support a wider

variety of organisms and are scored higher than a pool substrate dominated by

mud or bedrock and no plants.

• Velocity/Depth Regime (riffle/run)

Evaluates the presence/absence of four velocity/depth regimes (fast-deep, fast-

shallow, slow-deep, and slow-shallow). Generally, shallow is < 0.5m and slow is

< 0.3m/sec.

• Pool Variability (low gradient)

Evaluates the overall mixture of pool types found in streams, according to size

and depth (large-shallow, large-deep, small-shallow, and small-deep).

The next four parameters evaluate a larger area surrounding the sampled reach. This

expanded area is the stream length defined by at least how far upstream the

investigator can see from the downstream point of the sample reach, but at least 100

meters upstream of the sampled reach.

• Channel Alteration (riffle/run & low gradient)

Evaluates the extent of channelization or dredging, but can include any other

large-scale changes in the shape of the stream channel that would be

detrimental to the habitat. Channel alteration is present when artificial

embankments, riprap, and other forms of artificial bank stabilization or structures

are present; when the stream is very straight for significant distances; when

dams and bridges are present; and when other such changes have occurred.

• Sediment Deposition (riffle/run & low gradient)

Estimates the extent of sediment effects in the formation of islands, point bars,

and pool deposition. Deposition is typically evident in areas that are obstructed

by natural or manmade debris and areas where the stream flow decreases, such

as bends.

• Riffle Frequency (riffle/run)

Estimates the frequency of riffle occurrence based on stream width and thus the

heterogeneity occurring in a stream. For riffle/run prevalent streams where

distinct riffles are uncommon, a run/bend ratio is used as a measure of

meandering or sinuosity.

• Channel Flow Status (riffle/run & low gradient)

5-6

Estimates the areal extent of exposed substrates due to water level or flow

conditions. The flow status will change as the channel enlarges (e.g., aggrading

stream beds with actively widening channels) or as flow decreases as a result of

dams and other obstructions, diversions for irrigation, or drought. In riffle/run

prevalent streams, riffles and cobble substrate are exposed; in low gradient

streams, the decrease in water level exposes logs and snags, thereby reducing

the areas of good habitat.

The last four parameters apply to riffle/run prevalent as well as low gradient streams,

and evaluate an even greater area. This area is usually defined as the length of stream

that was electrofished (or an approximate 100-meter stream reach when no fish were

sampled). It can also take into consideration upstream land-use activities in the

watershed.

• Condition of Banks (riffle/run & low gradient)

Evaluates the extent of bank failure, signs of erosion, or the potential for erosion.

The stream bank is defined as the area from the water’s surface to the bankfull

delineation. Steep banks are more likely to collapse and suffer from erosion than

are gently sloping banks, and are therefore considered to be unstable. Signs of

erosion include crumbling, unvegetated banks, exposed tree roots, and exposed

soil.

• Bank Vegetative Protection (riffle/run & low gradient)

Estimates the extent of stream bank that is covered by plant growth providing

stability through well-developed root systems. The stream bank is defined as the

area from the water’s surface to the bankfull delineation. This parameter supplies

information on the ability of the bank to resist erosion as well as some additional

information on the uptake of nutrients by the plants, the control of instream

scouring, and stream shading. This parameter is made more effective by defining

the native vegetation for the region and stream type (i.e., shrubs, trees, etc.). In

some regions, the introduction of exotics has virtually replaced all native

vegetation. The value of exotic vegetation to the quality of the habitat structure

and contribution to the stream ecosystem must be considered in this parameter.

In areas of high grazing pressure from livestock or where residential and urban

development activities disrupt the riparian zone, the growth of a natural plant

community is impeded and can extend to the bank vegetative protection zone.

• Grazing or Other Disruptive Pressures (riffle/run & low gradient)

Evaluates disruptions to surrounding land vegetation due to common human

activities, such as crop harvesting, lawn care, excavations, fill, construction

projects, and other intrusive activities.

• Riparian Vegetative Zone Width (riffle/run & low gradient)

5-7

Estimates the width of natural vegetation from the edge of the stream bank out

through the riparian zone. Narrow riparian zones occur when roads, parking lots,

fields, lawns, bare soil, rocks, or buildings are near the stream bank. Residential

developments, urban centers, golf courses, and rangeland are the common

causes of anthropogenic degradation of the riparian zone. Conversely, the

presence of "old field" (i.e., a previously developed field not currently in use),

paths, and walkways in an otherwise undisturbed riparian zone may be judged to

be inconsequential to altering the riparian zone and may be given relatively high

scores.

After all parameters in the matrix are evaluated, the scores are summed to derive a total

habitat score for that station. Please refer to the Assessment Book, Chapter 4.1,

Physical Habitat Assessment Method (Walters 2017) for accompanying assessment

method.

LITERATURE CITED

Barbour, M. T., J. Gerritsen, B.D. Snyder, and J. B. Stribling. 1999. Rapid

bioassessment protocols for use in streams and wadeable rivers: periphyton,

benthic macroinvertebrates and fish, second edition. EPA 841-B-99-002. U.S.





Pennsylvania.

5-8

5.2 PEBBLE COUNT DATA COLLECTION PROTOCOL

5-9

Prepared by:

Gary Walters






2009

Edited by:

Gary Walters






2017

5-10

INTRODUCTION

This survey protocol is to be applied to riffle/run dominated, gravel or cobble bed stream

segments to determine substrate stability. Flow regime alteration (change in volume

and/or timing of discharge) is a major cause of stream instability and habitat alteration.

One aspect of concern is the delivery of fine sediments to streams and their effects on

aquatic habitat. One approach to monitoring these sediment effects is “A Pebble Count

Procedure for Assessing Watershed Cumulative Effects” by Bevenger and King (1995).

This procedure utilizes a reference stream approach in evaluating the stability of study

or candidate streams. The procedure characterizes particle size distributions of

reference and study streams, where reference streams are defined as “natural” or “least

disturbed” and study streams as “disturbed” or “impacted”. These particle size

distributions can be used for comparative purposes to determine, with statistical

reliability, if there has been a shift toward finer size materials in the study stream. This

protocol employs a modification of the Wolman (1954) pebble count procedure to a

zigzag pattern through a continuum along a longitudinal reach of the stream. This allows

for numerous meander bends and associated habitat features to be sampled as an

integrated unit.


Wadeable reference and study streams should be selected from the same ecoregion,

and the streams should be classified according to the Rosgen stream classification

system (Rosgen 1994, 1996) prior to conducting the field collection. Streams

classification can be accomplished in the office using topographic quadrangles and

aerial photographs, and the classification should be confirmed when the sample site is

visited. This protocol should only be applied to those streams that are classified as

B and C types with cobble (B3 or C3) or gravel beds (B4 or C4). If the classification

results in stream types G, F, or D, then data collection may not be necessary since, in

most cases, these stream types are the result of channel instability. If the instability

were a result of natural conditions the stream would not be classified as impaired. Also,

if the classification results in stream types A and E, which are ordinarily stable, then

data collection is not necessary. In addition, this procedure should not be conducted on

“natural” sand or silt/clay bottom streams, as fine particles will be the predominate

substrate type, thus resulting in potentially misleading indications of instability.

PARTICLE COUNT PROCEDURE

Once reference and study streams have been identified, the sample stream reach should

include at least two riffle and two pool habitat units if present, or a minimum of 200 meters.

5-11

The chosen sample reach habitat units should be representative of the streams. Study

and reference streams must have a minimum mean width of 3 meters. If mean stream

width is greater than 20 meters, then sample reach should be extended 100 meters for

each 10-meter increment increase in width. Sampling of reference streams should occur

within a few days of the sampling of study streams when possible and should always

occur within the same year and season. In order to confirm stream classification, at least

two stream cross-sections (one riffle and one pool) should be measured from bankfull

elevation to bankfull elevation within the study reach, prior to conducting the pebble count.

Pebble counts are conducted on the selected reach beginning at the head of a riffle and

continuing through 4 habitat units (2 riffle, 2 pools if present), or for a minimum of

200 meters. At least 200 particles are to be sampled from the stream reach. Pebble

counts are conducted along a zigzag transect from bank toe to bank toe in the active

channel (Figure 1). The angle of the transect from the bank should be maintained as

best as possible and can be aided by identifying a location to walk to on the opposite

bank. Particles are selected beginning at the start point by placing a finger at the toe of

one boot, and without looking, sliding a finger down to the stream bottom until making

contact with a particle (Figure 1). Each particle selected is measured along the

intermediate axis (Figure 1) and the measurement is recorded on the Pebble Count

Data Collection Form (Appendix C.4). Alternatively, each particle measurement may be

tallied according to Wentworth size classes (<2 mm, 2-4 mm, >4-8 mm, >8-16 mm, etc.)

on the Alternative Pebble Count Field Data Collection Form (Appendix C.5). The

investigator then paces off a chosen distance to the next point and samples another

particle in the same manner as the first. The distance to the next sample point should

be no less than 2.1 meters to avoid correlation between particles sampled.

Data analysis is conducted as described in the Assessment Book, Chapter 4.1, Physical

Habitat Assessment Method (Walters 2017) for habitat assessment method.

5-12

Figure 1. Zig-zag pebble count procedure from Bevenger and King, 1995.

5-13

LITERATURE CITED

Bevenger, G. S., and R. M. King. 1995. A pebble count procedure for assessing

watershed cumulative effects. U.S. Department of Agriculture, Fort Collins

Colorado.

Rosgen, David L. 1994. A stream classification system. Catena. Elsevier Science,

Amsterdam 22: 169–199.

Rosgen, David L. 1996. Applied river morphology. Wildlands Hydrology Books, Pagosa

Springs, Colorado.




Pennsylvania.

Wolman, M. G. 1954. A method of sampling coarse river-bed material. Transactions

American Geophysical Union 35(6):951–956.

5-14

5.3 WATER FLOW DATA COLLECTION PROTOCOL

5-15

Prepared by:

Josh Lookenbill






2009

Edited by:

Josh Lookenbill






2017

5-16

INTRODUCTION

Water flow (hereto referred to as discharge) is the volume of water that passes through

a conduit over a period of time. For the purposes of this document, the conduit is

typically an open stream channel, but it could also include storm drain infrastructure and

other water conveyances. Discharge data is collected along with other water quality

data in order to characterize yields or quantities of pollutants that are transported by

waterbodies. Chemical data is collected and reported as concentrations. Discharge data

allows those concentrations to be further characterized as standardized quantities or

yields. This is required in characterizing the amount of a pollutant transported by a

waterbody that may be affecting water quality.

Historically, DEP has collected discharge data by implementing cross-section

measurements using the “Mid-Section” approach described by Buchanan and Somers

(1969); where depth and velocity is measured at subdivisions across a transect,

discharge is calculated at each subdivision, and the discharge at each of the

subdivisions is summed to calculate total discharge. This approach has been utilized in

small to medium wadable waterbodies and continues to be a valid discharge data

collection protocol.

Over time the utility and consequently the number and frequency of discharge

measurements has dramatically increased. Routine discharge is required to regulate

water allocation, pollution control programs, generate flood prediction forecasts, and

support other important socioeconomic functions. The USGS originally set the standard

for collecting discharge data with USGS Techniques of Water Resources Investigations

book 3, chapter A8 (TWRI 3–A8), “Discharge Measurements at Gaging Stations” by

Buchanan and Somers (1969). Since then technology and data collection protocols

have evolved, and USGS has continued to set the standard for collecting discharge

data. Additional USGS publications include “Measurement and Computation of

Streamflow, volumes 1 and 2,” by Rantz et al. (1982); “Discharge measurements using

a broad-band acoustic Doppler current profiler,” by Simpson (2002); “Quality-assurance

plan for discharge measurements using acoustic Doppler current profiler,” by Oberg et

al. (2005); and “Measuring discharge with acoustic Doppler current profilers from a

moving boat,” by Mueller and Wagner (2009).

DEP no longer maintains a detailed discharge data collection protocol, but references

and implements the latest USGS techniques and standards for collecting discharge

data. USGS Techniques and Methods book 3, chap. A8, “Discharge Measurements at

Gaging Stations” (Turnipseed and Sauer 2010) is the latest techniques and standards

that supersedes or supplements previous USGS publications.

5-17

LITERATURE CITED

Buchanan, T. J., and W. P. Somers. 1969. Discharge measurements at gaging stations.

Book 3, chapter A8, page 65. in National field manual for the collection of water

quality data: Techniques and methods. U.S. Department of the Interior, U.S.

Geological Survey, Reston, VA. (Available online at:

https://pubs.er.usgs.gov/publication/tm3A8.)

Mueller, D. S. and C. R. Wagner. 2009. Measuring discharge with acoustic Doppler

current profilers from a moving boat. Book 3, section A, chapter 22. in National

field manual for the collection of water quality data: Techniques and methods.

U.S. Department of the Interior, U.S. Geological Survey, Reston, VA. (Available

online at: https://pubs.usgs.gov/tm/3a22/.)

Oberg, K. A., S. E. Morlock, and W. S. Caldwell. 2005. Quality assurance plan for

discharge measurements using acoustic Doppler current profilers: United States

Geological Survey Scientific Investigations report 2005–5183. U.S. Department

of the Interior, U.S. Geological Survey, Reston, VA. (Available online at:

https://pubs.usgs.gov/sir/2005/5183/SIR 2005-5183.pdf.)

Rantz, S. E. 1982. Measurement and computation of streamflow: Water supply paper

2175. U.S. Department of the Interior, U.S. Geological Survey, Reston, VA.

(Available online at: https://pubs.er.usgs.gov/publication/wsp2175.)

Simpson, M. R. 2002. Discharge measurements using a Broad-Band Acoustic Doppler

Current Profiler: United States Geological Survey Open-File Report 01–01. U.S.

Department of the Interior, U.S. Geological Survey, Sacramento, CA. (Available

online at: https://pubs.usgs.gov/of/2001/ofr0101/.)

Turnipseed, D. P. and V. B. Sauer. 2010. Discharge measurements at gaging stations:

United States Geologic Survey. Book 3, chapter A8. in National field manual for

the collection of water quality data: Techniques and methods. U.S. Department of

the Interior, U.S. Geological Survey, Reston, VA. (Available online at:

https://pubs.usgs.gov/twri/twri3a8/.)

A-1

APPENDIX A: BIOLOGICAL DATA COLLECTION INFORMATION AND FORMS

A-2

A.1 MACROINVERTEBRATE FIELD DATA SHEET

A-5

A.2 MACROINVERTEBRATE ENUMERATION BENCH SHEET

A-7

A.3 FISH FIELD DATA COLLECTION SHEET

B-8

FIELD DATA SHEET SEMI QUANTITATIVE FISH SURVEY SAMPLING

Site ID: Stream Name: Date:

Crew/Agency:

Location:

Reach Desc:

County: Municipality:

Start-End Time (Military): -

Mean Site Width (M): Site Length (M):

Latitude (dec. deg.): Longitude (dec. deg.):

Temperature (°C):

Sp. Conductance (µs/cmc):

pH (SU):

DO (MG/L):

Alkalinity (MG/L):

Habitat Score:

Water Clarity: Turbidity (NTU): Secchi Depth: Meters

Or if you forgot your Secchi Disk or Turbidity Meter circle one: 1 2 3 4 5 6 7 8 9 10

(Turbid ≤ 1 m) (Discolored 1-2 m) (Clear > 2 m)

Gear(s):

Backpack AC Volts: Button time seconds: # passes:

Towboat DC Watts: Pulse rate (Hz): # netters:

Boat P-DC Amps: Pulse Width (ms): # probes:

Make/Model:

Comments:

% Mesohabitat:

Riffle % Run % Pool %

% Velocity/Depth Regimes

Slow-Deep % Slow-Shallow % Fast-Deep % Fast-Shallow %

% Habitat: (% fish in habitat (# individuals) / % habitat within reach)

Root Wads / % Mud Bank __/__% Macrophyte / % Artificial (Docks, Piers, etc.) / %

Boulder / % Cobble / % Gravel / % Sand / % Silt / %

Large Woody Debris / % Small Wood Debris / %

Comments:

% Substrate:

Boulder % Cobble % Gravel % Sand % Silt %

Detritus % Bedrock % Artificial or Channelized %

Comments:

A-11

FIELD DATA SHEET SEMI-QUANTITATIVE FISH SURVEY SAMPLING (continued)

Site ID:

Stream Name: Date:

Abv. Common

Name DELTP* Description BODY

Bg Black grub (Black spot) P Small black cyst approximately one millimeter in diameter, caused by a larval trematode.

Pe External parasite P Ectoparasite attached to body, fins or gills (Leech, Fish louse, Anchor parasite etc.) De Deformities D Body structure is abnormal, ex. curvature of the spine Fi Fungal infection L Fungal infections may include a fuzzy (cotton) appearance w/ discolored areas or

lesions. Ma Melanistic area L Darkly pigmented spot(s), not raised from the skin. Os Open sore L Lesions, (Note any mucus or necrotic tissue that may be in or around the sore) Rr Raised red sore L A bulging or bubbled red sore Sh Scales hemorrhagic L Evidence of bleeding or hemorrhaging at base of scales Tu Tumor T Unusual mass or fatty growth Cw White cysts P Usually a small crème-colored wart or bubble appearance Cc Caudal cysts P Small cysts specifically found on caudal peduncle, or at the base of anal/dorsal

fins Em Emaciated D Body thin and lacks normal robustness, "starved"

EYES Ec Eye cloudy L Eye cloudy or opaque Ee Eye exophthalmic L Eye bulging out of socket Eh Eye hemorrhagic L Eye bleeding or has evidence of burst blood vessels Ed Eye deteriorated L Eye deteriorated, obviously blind, eye missing completely

FINS Fe Fins eroded E Erosion of the fin (make note of spawning, as some species will erode fins during

this active time) Fh Fins hemorrhagic E Evidence of bleeding or hemorrhaging of fins Ff Fins frayed E Fins appear worn out and stressed (make note of spawning)

GILLS Ge Gills eroded E Necrotic tissue or eroded to the point that sections of the gill filaments are

missing Gf Gills frayed E Gill lamellae appear worn out and stressed Gs Gill spots P Small white spots throughout the lamellae Gc Gill cysts P Cyst attached to gills, usually a small crème-colored wart or bubble appearance

OTHER G Gravid

Female fish full of eggs, releases upon handling

P Post spawn

Egg sack has already released, abdominal wall stretched but empty, (Only if OBVIOUS)

M Milting

Male fish releases milt upon handling

Record as- Species- Length(to nearest 5 mm) Anomaly

Example- Smallmouth bass- 310WcEc, 470, 225Ge *DELTP- Deformities, Eroded fins/gills, Lesions, Tumors, Parasites

A-12



SPECIES (length > 25mm) #

Photo(s) Field Count

Lab Count Total Count

1.

2.

3.

4.

5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

A-13



SPECIES (length > 25mm) #

Photo(s) Field Count

Lab Count Total Count

30.

31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

Comments:

A-14

GENERAL INSTRUCTIONS FOR FIELD DATA SHEET SEMI-QUANTITATIVE FISH SURVEY SAMPLING

Station ID: YEARMMDD-TIME-AGENCY, example: (20080601-1200-DEP). TIME = START TIME

Crew/Agency: List each crew member and agency or affiliation.

Location: General description, example: Transect is located 500 meters north of Pine Run and 1000 meters south of Rt. 441 bridge.

Start-End Time: Enter military time, start of shocking to end of shocking.

Mean Site Width: In the first 100 meters, 5 wetted channel widths are measured using a graduated measuring tape or range finder (every 20 meters from the starting point) and averaged.

Site Length: Wadeable = Minimum site length of 100 meters should be surveyed. Increased site length can be used as necessary to cover all habitats (pools, riffles, runs, and cascades).

Average Stream Width (m) Minimum Site Length

<10m 100m

10 to 40m 10 times the average stream width

>40m Maximum of 400m

Nonwadeable = Minimum site length of 500 meters should be surveyed. Increased site length can be used as necessary to cover all habitats (pools, riffles, runs, and cascades).

Latitude/Longitude: Decimal Degrees.

Current: Check one.

Species: List species and count all individuals > 25mm.

Column – Photo #: Number fish photographed.

Column – Field Count: The number of the fish identified in the field.

Column – Lab Count: The number of the fish identified in the laboratory.

Comments: Water/weather conditions, problems, etc.

A-15

A.4 FISH TISSUE FIELD DATA COLLECTION SHEET

A-16

COMMONWEALTH OF PENNSYLVANIA

DEPARTMENT OF ENVIRONMENTAL PROTECTION

BUREAU OF WATER STANDARDS AND FACILITY REGULATION

GENERAL INSTRUCTIONS FOR

FIELD DATA SHEET TISSUE SAMPLING

STATION Enter WQN Station Number, WQF Station Number or Survey Station Number as

NUMBER: appropriate. Leave blank if none of these apply.

NOTE: If WQN Number is entered, then Waterbody Name, Location, County and Municipality can

be left blank.

DATE, COLLECTOR, AGENCY and COLLECTOR NUMBER: Must be completed.

PROJECT ID: FISH

NEW STATIONS: If a WQF station number has not previously been assigned, Quad. Name, Quad. Number, either

Lat/Long or inches-N/inches-W and RMI (to nearest 0.1 mile) must be entered (for STORET input).

TISSUE TYPE: Standard samples are:

1. Skin-on, scaled fillets for gamefish, panfish and rough fish.

2. Skinless fillets for catfishes.

3. Skinless one-inch cross-sections for American Eel

SAMPLE: A normal sample is a composite of 10 fillets from 5 fish. For large fish, 5 fillets can be used.

75 PERCENT RULE: The individual fish in each composite should be as close to the same size as possible. A general

guideline is that the length of the smallest individual in the composite should be no less than 75

percent of the length of the largest individual.

Black grub (Black spot) Bg Tumor Tu Fins eroded Fe

External parasite Pe White cysts Cw Fins hemorrhagic Fh

Deformities De Caudal cysts Cc Fins frayed Ff

Fungal infection Fi Emaciated Em Gills eroded Ge

Melanistic area Ma Eye cloudy Ec Gills frayed Gf

Open sore Os Eye exophthalmic Ee Gill spots Gs

Raised red sore Rr Eye hemorrhagic Eh Gill cysts Gc

Scales hemorrhagic Sh Eye deteriorated Ed

Gravid G Post spawn Ps Milting male M

Fish Health:

Life History:

A-17

COMMONWEALTH OF PENNSYLVANIA

DEPARTMENT OF ENVIRONMENTAL PROTECTION

BUREAU OF WATER STANDARDS AND FACILITY REGULATION

FIELD DATA SHEET TISSUE SAMPLING

Station # Waterbody Date & Time

Location

County Municipality

Collector Agency Coll.#

COMPLETE FOR NEW STATIONS

Quad. Name Quad.#

Lat. (inches N) Long. (inches W) RMI

Protocol: Electrofishing

Seine

Rotenone

Angling

Tissue Type: Whole Fish

Skinless Fillet

Skin-on Fillet – Scaled

Skin-on Fillet – Not Scaled

Other (specify):

SPECIES TL (inches) WT (lbs. oz.) Fish Health & Life History*

1.

2.

3.

4.

5.

6.

7.

8.

9.

10.

*Use Fish Health & Life History Codes (if needed)

Comments: (water/weather conditions, man-hours expended, problems, etc.)

A-18

A.5 MUSSEL FIELD DATA COLLECTION SHEETS

A-19

Date: Site#: Location:

Water Temperature: Air Temperature: page of

Dive #

Distance

from X site

Distance

from Shore Latitude Longitude Diver

Backup

Diver Air In Air Out Time In

Time

Out

Invasive

Mussel Score

and Species

Max

Depth

Comments: Bucket Dive

% Boulder: % Boulder:

% Cobble: % Cobble:

%Gravel: %Gravel:

%Sand: %Sand:

%Silt: %Silt:

Dive #

Distance

from X site

Distance


Backup


Time

Out

Invasive

Mussel Score

and Species

Max

Depth



% Cobble: % Cobble:

%Gravel: %Gravel:

%Sand: %Sand:

%Silt: %Silt:

Dive #

Distance

from X site

Distance


Backup


Time

Out

Invasive

Mussel Score

and Species

Max

Depth



% Cobble: % Cobble:

%Gravel: %Gravel:

%Sand: %Sand:

%Silt: %Silt:

A-20

Date: Location:

Site# Dive# Bag# Genus Species

Live or

Dead

Length

(mm)

Width

(mm)

Depth

(mm)

Weight

(g)

Weight W/O

Invasive

Mussel

Attached

Invasive

Mussel

Length

(mm) Notes

A-21

A.6 QBE PERIPHYTON FIELD DATA COLLECTION SHEET

A-22

Time(UTC-5)

Temp (°C)

SPC(µS/cm)

> 10 in. pH

2.5-10 in. DO (mg/L)

0.1 - 2.5 in DO (%)

0.06-2 mm Turb (NTU)

0.004-0.06

<0.004 mm

All

> 25 meters

> 25 meters

All Quantity

All Rocks

> 25 meters Sampled

> 25 meters

All

Transect 1

RDB LDB W / 1/3

Transect 2

RDB LDB W / 1/3

Transect 3

RDB LDB W / 1/3

Transect Random Rock Pick Calculation (Watershed area ≥ 1,000 sq mi = 27-rock & area < 1,000 sq mi = 9-rock)

Random #'s: Random #'s: Random #'s:

Random #'s: Random #'s: Random #'s:

3/4 Downstream

Left Descending Bank

Reach Orientation: DWS Facing Orientation taken at Densiometer 1/2

Downstream position (compass degrees CW from magnetic north)

Shaded Dots

Densiometer

Reading/Orientation

Right Descending Bank

1/4 Upstream

Canopy Closure Canopy Closure

Leaf-Off Leaf-On

(Middle)

(Downstream) Random #'s: Random #'s: Random #'s:

Sand

PADEP QBE (Quantitative Benthic Epilithic) Periphyton Field Data Sheet - Page 1

Record at

stream

widths

% Canopy Closure/Density = Divide total by 68 (< 25m) or Divide by 136 (>25m)

(Upstream)

Total / % Closure

1/2 Upstream

1/2 Downstream

3/4 Upstream

Shaded Dots

Inorganic Substrate: % relative

Station ID

Chem Coll-Seq

Date

Time - Arrival

Personnel

1/4 Downstream

Silt

Clay

Bedrock

Boulder

Cobble

Gravel

Latitude

Longitude

Days Stable flow

Site / Sample Information

Watershed Area (mi2)

Chem SAC Code

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Collector # Sequence #



PADEP QBE - 2 (Quantitative Benthic Epilithic) Periphyton Field Data Sheet - Page 2

Sub-Sample Processing Information:

AFDM Filter & Notes - 2ml on GFF filter - foil wrapped. (Collect 3 replicate filters).

Store/transport on dry ice then store at -80°C in BOL Bacti freezer.

Phycocyanin Bottle & Notes - 30ml in 120ml white plastic sample bottle.

Store/transport on wet ice. Upon arrival at BOL deliver to BOL Bacti refrigerator. BOL

will pull 3 sub-samples from this single bottle.

Algal ID Bottles & Notes - 100ml well homoginized slurry in 120ml white plastic sample

bottle + 7ml formaldehyde preservative (Collelct min three 100ml replicate bottles).

Total Diluted Algae Slurry Volume (ml): Dilute as needed to achieve

min required: 700ml with cyanotoxin test, 400ml without cyanotoxin

If col lecting for cyanotoxin testing: 250ml in whirl -pak bag then place whirl -pak in ziplock bag. Store/transport

on dry ice then s tore at -80°C in Monitoring Section freezer.

Chlorophyll-a Filter & Notes - 2ml on GFF filter with 1ml MgCO3 fixative - foil wrapped.

(Collect 3 replicate filters). Store/transport on dry ice then store at -80 °C in BOL Bacti

freezer. Indicate if submitting for acidified or non-acidified processing.

Other Notes

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A.7 QMH PERIPHYTON FIELD DATA COLLECTION SHEET

A-25

Time(UTC-5)

Temp (°C)

SPC(µS/cm)

pH

DO (mg/L)

DO (%)Turb (NTU)

All

> 25 meters

> 25 meters

All

All

> 25 meters

> 25 meters

All

1/2 Upstream

Right Descending Area

1/4 Upstream

1/4 Downstream

Stream Order

Total Diluted Algae Slurry Volume

(ml): Dilute as needed to achieve

min required: 700ml with

cyanotoxin test, 400ml without

cyanotoxin

Personnel

PADEP QMH (Qualitative Multi-Habitat) Periphyton Field Data Sheet - Page 1

Station ID Latitude

Site / Sample Information

Date

Time - Arrival Days Stable flow

Chem Coll - Seq

Chem SAC Code Longitude

Other % Other %

Periphyton Habitats: % relative qualification

Epidendric (Wood) %

Epiphytic (Plant) % a

Epilithic (Rock) %

Episammic (Sand) %

Epipelic (Silt) % Gravel %

Reach Orientation: DWS Facing Orientation taken at Densiometer 1/2

Downstream position (compass degrees CW from magnetic north)

Densiometer

Reading/Orientation

Canopy Closure Canopy Closure Record at

stream

widths

Leaf-Off Leaf-On

Shaded Dots Shaded Dots

1/2 Downstream

3/4 Upstream

Total / % Closure

% Canopy Closure/Density = Divide total by 68 (< 25m) or Divide by 136 (>25m)

3/4 Downstream

Left Descending Area

A-26

Other Notes: a Record -plant taxa from which sampled col lected. If collecting for cyanotoxin testing: 250ml in whirl -pak

bag then place whirl -pak in ziplock bag. Store/transport on dry ice then s tore at -80°C in Monitoring Section freezer.

Algal ID Bottles & Notes - 100ml well homoginized slurry in 120ml white plastic sample bottle + 7ml

formaldehyde preservative (Collect min three 100ml replicate bottles).

PADEP QMH (Qualitative Multi-Habitat) Periphyton Field Data Sheet - Page 2

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A.8 ALGAE IDENTIFICATION AND ENUMERATION LABORATORY SUBMISSION DATA

SHEET

A-28

PA DEP GIS Key

IDDate

Sample Type

(QMH = Quitative

Multi-Habitat, QBE

= Quantitative

Benthic Epilithic)

Stream Name Location Description County Latitude Longitude

Surface

Area

(cm^2)

Initial Sample

Volume (ml)

ID Bottle

Sub-

Sample

Volume

(ml)

Formaldehyde

Added (ml)

Algae Identification and Enumeration Laboratory Submission Data Sheet

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A.9 MICROBIOLOGY SAMPLE SUBMISSION SHEET FOR ACIDIFIED CHLOROPHYLL-A,

PHYCOCYANIN, AND AFDM

A-31

A.10 BACTERIA MONITORING FIELD AND SITE DATA SHEET

B-1

APPENDIX B: CHEMICAL DATA COLLECTION INFORMATION AND FORMS

B-2

B.1 FIELD METER CALIBRATION FORM

B-4

B.2 TRANSECT DATA COLLECTION FORM

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B.3 SAMPLE INFORMATION SYSTEM USER’S GUIDE

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o Formalized classroom training o Small group training o One-on-one training

• Participate in meetings to provide application guidance

• Telephone support help desk

• Application web page development and maintenance

• Publish articles identifying solutions to common problems

• Application testing

• Documentation development

• Application on-line help development and maintenance

WHAT IS NHD?

The National Hydrography Dataset (NHD) is a set of digital spatial data created by the USGS

and USEPA containing information about naturally occurring and constructed bodies of water,

natural and artificial paths through which water flows, and related hydrographic entities. These

entities, or features, are combined to form reaches, which provide the framework for linking (or

geo-coding) water-related data to the NHD surface water drainage network.

ACCESSING SIS

Security

Security roles assigned to a username determines a user’s ability to create, update, or delete

information in SIS. To add or modify security roles, contact a system coordinator to submit a

security request. The available security roles are:

• Sample Query

• Sample Entry

• Project Entry

Logging on to SIS

A user with security access to SIS can login as follows:

1. Open the IntraDEP page (http://intradep/). 2. Click the “Oracle Applications” link. 3. Click the “SIS – Samples Information System” link and the Logon pop-up window

displays.

The first time accessing Oracle applications several security prompts display. Reference the “Special note regarding security warnings” located at the top of the “Oracle Applications” page for instructions on responding to the security prompts.

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Figure 1: The Logon pop-up window.

4. Click in the Username field and enter a user name. 5. Click in the Password field and enter the password. 6. Click in the Database field and enter “PROD”. 7. Click the CONNECT button to log on to SIS.

Changing an Oracle Password

Each user has a single password for all Oracle applications. Changing the password used to

access SIS also changes the password for eFACTS, CTS, etc.

Change an Oracle password as follows:

1. Open the IntraDEP page (http://intradep/). 2. Click the “Oracle Applications” link. 3. Click the CHANGE PASSWORD button and the Logon pop-up window displays. 4. Click in the Username field and enter a user name. 5. Click in the Password field and enter the current password. 6. Click in the Database field and enter PROD. 7. Click the CONNECT button and the “Change Oracle Password” pop-up window

displays.

Figure 2: The “Change Oracle Password” pop-up window.

8. Click in the Old Password field and enter the current password. 9. Click in the New Password field and enter a new password. 10. Click in the Confirm Password field and enter the new password a second time. 11. Click the OK button. 12. A message in the hint line at the bottom of the screen will indicate if the password was

successfully changed.

SIS OVERVIEW

This section provides an overview of the screen layout, common structure, fields, and buttons

within SIS.

Title Bar

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The Title Bar is located at the top of each screen in SIS and displays the name of the screen.

The Title Bar also includes minimize, maximize, and close window buttons that change the

size of the active window.

Menu Bar

The Menu Bar is located below the Title Bar and displays two different layers: the Main Menu

Bar and the Screen Menu Bar.

Main Menu Bar

The Main Menu Bar is only available on the SAMPLE INFORMATION SYSTEM Screen that

displays after first logging in to SIS. The Main Menu Bar includes menus for each screen in

SIS.

Screen Menu Bar

The Screen Menu Bar is available on each screen in SIS and contains menus, commands, and

sub-commands to access reports, functions, and other screens in SIS.

The Screen Menu Bar has two levels: menus and commands. The first-level menus are

always visible at the top of the screen and do not directly access a screen or report. Click a

menu to display a list of commands. Click a command to perform an action such as displaying

a screen, saving a record, printing, or displaying a second-level of commands.

Toolbar

Every screen in SIS contains a horizontal toolbar positioned at the top of the window under the

Screen Menu Bar. The toolbar displays in two modes: Entry Mode and Query Mode.

Entry Mode

The Entry Mode toolbar displays by default after opening a screen.

Save (F10)

The SAVE button commits to the database any created records or updates and

deletions of existing records on the active screen. Information entered on a screen is not

added to the database until the modifications are committed to the database by clicking the

SAVE button.

Print

The PRINT button sends the visible information displayed on the screen to a printer.

Clicking the PRINT button displays a pop-up window to select a printer, paper size, print

orientation, and number of copies.

Enter Query (F7)

The ENTER QUERY button displays when the active screen is in Entry Mode. Clicking

the ENTER QUERY button changes the screen from the Entry Mode to Query Mode, which is

used to search for records in SIS. Clicking the ENTER QUERY button replaces it with three

Query Mode buttons: EXECUTE QUERY, CANCEL QUERY, and COUNT HITS. For more

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information on the three Query Mode buttons, reference the QUERY MODE section of the user

guide.

Next Record ( )

The NEXT RECORD button displays, or navigates to, the next record of the current

block. Use this button to view additional records when a query is executed and retrieves more

than one record.

Previous Record ( )

The PREVIOUS RECORD button displays, or navigates to, the previous record of the

current block.

Create Record (F6)

The CREATE RECORD button clears the current record from the screen and creates a

new, blank record.

Delete Record (Shift + F6)

The DELETE RECORD button deletes the current record. Clicking the DELETE

RECORD button will delete the current record on which the cursor is residing. The record is

not permanently deleted until the SAVE button is clicked.

Duplicate Record

The DUPLICATE RECORD button duplicates the previously displayed record into the

current, empty record. Update any fields on the duplicated record and then save the changes.

Use this button to enter multiple similar records.

Edit Item (Ctrl + E)

The EDIT ITEM button displays the Editor pop-up window with the contents of the

currently selected field. The Editor pop-up window provides the user with a larger space to

view or update the contents of a field.

Clear Record (Shift + F4)

The CLEAR RECORD button clears the current record from the screen without deleting

it. This button removes the currently displayed record from the buffer of queried records and

displays the next record in the queried buffer.

Clear Form

The CLEAR FORM button clears the current screen of all records without updating the

records. Clicking the CLEAR FORM button places the screen in a similar state to when it is

first opened.

Help

The HELP button provides the technical properties of the field in which the cursor

resides.

Exit (CTRL + Q)

The EXIT button closes the current screen.

QUERY MODE

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Use Query Mode to display records that match user-specified criteria. Screens open in Entry

Mode with the ENTER QUERY button visible. Click the ENTER QUERY button to place the

screen into Query Mode, which hides the ENTER QUERY button and displays the EXECUTE

QUERY, COUNT HITS and CANCEL QUERY buttons.

Execute Query (F8)

The EXECUTE QUERY button retrieves all records that match the user-defined query

criteria.

Caution: If no criteria are specified, a “blind query” is executed which retrieves all records.

Running blind queries is not recommended.

Count Hits (Shift + F2)

The COUNT HITS button displays a number in the hint line indicating how many records

match the query criteria. Click this button prior to executing a query to see the number of

results and determine whether to specify additional criteria to limit the results.

Cancel Query (Ctrl + Q)

The CANCEL QUERY button exits QUERY MODE and places the screen into the Entry

Mode. The CANCEL QUERY button does not stop an executed query that is currently

running.

SCREEN STRUCTURE

The screens in SIS use a standard layout with common elements described in this section.

Header and Detail Block

A header block identifies a specific sample displayed on the screen.

A detail block displays information related to the sample identified in the header block.

Figure 3: The SAMPLE ENTRY Screen with the header and detail blocks identified.

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Tabs

Detail blocks are separated into tabs that display additional information related to the record in

the header block. Click on a tab name to activate it and display the related details. The

header block continues to display the current sample record and the activated tab displays the

related detail record. The label of the currently displayed tab displays underlined and in red

font.

List of Values (F9)

A list of values (LOV) button, typically located to the right of a field, displays a list of codes

or a calendar for the related field.

Figure 4: A list of values with the first value selected.

Using the list of values button:

1. Click the list of values button to the right of the field and a list of values pop-up window displays.

2. Click on an entry in the list to select it.

The list may contain numerous values. Filter the list by clicking in the Find field, entering a wildcard (%) followed by part of a code or description, and then clicking the FIND button. The list displays only values containing the text entered in the Find field.

Click the OK button to select the highlighted entry or click the CANCEL button to return to the

screen without retrieving a value.

Pop-Up Messages

Pop-up messages alert or warn the user about certain actions, information, or functionality

throughout the system. Users should read the pop-up message for directions to follow or to

confirm an action.

Figure 5: A pop-up message confirming an action.

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Hint Line

Every window in SIS, except the SAMPLE INFORMATION SYSTEM Screen, contains a

horizontal bar at the bottom of the screen called the hint line. The hint line displays information

about the screen, an operation in progress, the result of an operation, or a description of the

currently selected field.

Figure 6: The hint line displaying a description for the Collector ID field.

QUERYING FOR SAMPLES

The following sections provide information on how to query an existing sample and the

associated details using the SAMPLES Screen. Access the SAMPLES Screen by clicking the

Samples Menu and then clicking the Query samples command.

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Figure 7: The samples screen.

Querying Using the Collector and Date Collected

This section provides steps for querying an existing sample using the collector, date collector,

and/or sequence number.

1. Access the SAMPLES Screen and then click the ENTER QUERY button on the toolbar or press the [F7] key.

2. Enter the four-digit collector ID assigned to the employee or monitoring point that collected the sample or select it by clicking the list of values button to the right of the Collector ID field. Press the [TAB] key.

3. If known, enter the date that the sample was collected. Press the [TAB] key. 4. If known, enter the time that the sample was collected. Press the [TAB] key. 5. If known, enter the sequence number assigned to the sample. 6. Click the EXECUTE QUERY button on the toolbar or press the [F8] key. 7. If more than one sample is retrieved, use the NEXT RECORD button on the toolbar or

press the [ ] key to locate the correct record.

If more than one record was retrieved, “Record: 1/?” displays below the hint line.

Querying using the Query Options Button

The Query Options button allows querying using a range of dates, sequence numbers, and

test results.

1. To use the QUERY OPTIONS button, access the SAMPLES Screen and then click the ENTER QUERY button on the toolbar or press the [F7] key.

2. Click the QUERY OPTIONS button and the “Query Options” pop-up window displays.

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Figure 8: The “Query Options” pop-up window.

3. Enter query criteria in any of the fields on the screen. 4. Click the COUNT HITS button on the toolbar to determine how many records will be

retrieved. 5. If the query count is extensive, further qualify the query before executing. 6. Click the EXECUTE QUERY button on the toolbar or press the [F8] key. 7. If more than one sample is retrieved, use the NEXT RECORD button on the toolbar or



8. To view the results and details of the sample, reference the VIEWING SAMPLE DETAILS section of the user guide.

Querying using Multiple Criteria

This section provides steps for querying a sample using the fields in the header block and the

fields under the detail tabs on the SAMPLES Screen.

If the complete query value for a field is not known, use the wildcard (%) at the beginning and/or end of one or two key characters. For example, to match both Lower Paxton and Paxtang municipalities enter: %PAXT%

1. Access the SAMPLES Screen and then click the ENTER QUERY button on the toolbar or press the [F7] key.

2. Click on a tab that contains a desired field for entering query criteria.

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Figure 9: The samples screen with the Name & Address TAB selected.

3. Enter query criteria in any of the queryable fields identified at the end of this section. 4. Click the COUNT HITS button on the toolbar to determine how many records will be

retrieved.

If the query count is extensive, further qualify the query before executing.

5. Click the EXECUTE QUERY button on the toolbar or press the [F8] key. 6. If more than one sample is retrieved, use the NEXT RECORD button on the toolbar or



7. To view the results and details of the sample, reference the Viewing the Sample Details section of the user guide.

Queryable Fields:

Sample Query Header Block

• Collector ID* • Date Collected

• Sequence# • Responsible Unit*

Project/Facility/Monitoring Pt TAB

• Project Unit* • Project ID*

• Primary Facility* • Sub-Facility*

• Monitoring Point ID#* • Monitoring Point Alias*

Name/Address TAB

• Addressee • Address Line 1

• Address Line 2 • City

• Zip Code • Phone#

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Sampling Conditions TAB

• Sampling Reason* • Sampling Protocol*

• Medium Type* • Medium*

• Stream Condition* • Initial Flow

• Final Flow • Units*

• Determination* • Depth To Water

• Units* • Determination*

Location TAB

• State* • County

• Municipality • Latitude

• Longitude • Datum

• Reference Point* • Geometric Type*

• Location Procedure* • Location

• Elevation • Accuracy

• Altitude Datum* • Method *

Comments/Quality Assurance TAB

• Appearance • Duplicate of Sample

• Quality Assurance Type* • Voided/Dry Sample

• Confidentiality Reason*

Administrative TAB

• Cost Center* • Entry Complete

• Date Created • Created By

• Date Last Modified • Last Modified By

*A list of values is available.

VIEWING SAMPLE DETAILS

This section provides steps for how to view the details associated with a sample.

1. Access the SAMPLES Screen and query a sample record. 2. Click the SHOW RESULTS button to display the SAMPLE ANALYSES/RESULTS

Screen, which includes the lab results of sample testing.

Figure 10: The SAMPLE ANALYSES/RESULTS Screen.

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3. The SAMPLE ANALYSES/RESULTS Screen displays the laboratory that tested the sample and a list of test codes with results. If there are multiple laboratory sample analyses, then click the NEXT RECORD or PREVIOUS RECORD buttons view a different analysis.

4. To view details about the laboratory identifiers for the sample, click the DETAILS button on the right-hand side of the screen. The “Lab Sample” pop-up window displays.

Figure 11: The “Lab Sample” pop-up window.

5. If available, click the LEGAL SEALS or COMMENTS button to view additional details. 6. Click the to exit the pop-up window and return to the SAMPLE

ANALYSES/RESULTS Screen. 7. To view the complete details of a specific test, click on a test code in the

Analyses/Results section of the screen to select it and then click the DETAILS button located at the bottom of the screen. The “Analysis/Result Details” pop-up window displays.

Figure 12: The “Analysis/Result Details” pop-up window.

8. If available, click the COMMENTS button to view additional comments. 9. Click the to exit the pop-up window and return to the SAMPLE

ANALYSES/RESULTS Screen. 10. Click the again to return to the SAMPLES Screen. 11. Six tabs containing information about the collection of the sample display in the detail

block of the screen.

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Figure 13: The six detail tabs on the samples screen.

12. Click on a tab to display the details:

• Project/Facility/Monitoring Pt TAB This tab identifies the specific project for which a sample was taken; the regulated entities (primary and sub-facilities) that may impact the soil, water, or air sample; and/or the monitoring point established for sample collection.

• Name & Address TAB This tab displays the name and address at which the sample was collected.

• Sampling Conditions TAB This tab displays the conditions under which the sample was collected including the sampling reason, method, medium, and stream condition.

• Location TAB This tab displays the location at which the sample was collected including county, municipality, latitude/longitude, and locational metadata.

• Comments/Quality Assurance TAB This tab displays comments and quality assurance indicators about the sample.

• Administrative TAB This tab displays cost center associated with the sample, who created and last updated the sample, and when the sample was created and last updated.

13. To generate a printed a report displaying the laboratory that tested the sample, a summary of the analysis methods, and results of the tests, click the PRINT REPORT button on the SAMPLE QUERY Screen.

EXPORTING A SAMPLE

The EXPORT button located on the SAMPLES Screen generates a customizable report using

the queried samples. The report is saved as file on the local computer. Contact the

Applications Helpdesk to request the “Sample Export” security role required to use this

function.

1. Access the SAMPLES Screen and query sample records to export. 2. Click the EXPORT button and the “Export Samples” pop-up window displays.

Figure 14: The “Export Samples” pop-up window.

3. Select which samples to include in the report and then click the CONTINUE button. The EXPORT SAMPLES Screen displays.

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Figure 15: The EXPORT SAMPLES Screen.

4. The column on the left-hand side of the screen lists items available to include in the report. To add an item, click on it and then click the > button.

5. The column on the right-hand side of the screen lists the items selected to include in the report. To remove an item, click on it and then click the < button.

Remove all selected items by clicking the << button or add all available items by clicking the >> button. To return to the selected items that initially display, click the “dflt” button.

6. Click the FORMAT OPTIONS button to specify how to delimit, or separate, the items selected for the report. Use the comma delimited option to generate reports that open in a spreadsheet application like Microsoft Excel or use the fixed width field option to generate reports to open in a word processing application like Microsoft Word.

After selecting items for export and setting a format, click the SAVE PROFILE button to save these settings to a profile on the local computer. Click the RETRIEVE PROFILE button to use the same settings when exporting a different set of samples.

7. Click the VIEW SAMPLE(S) button to verify the list of samples that the report will include.

8. Click the EXPORT DATA button and a save dialog displays. 9. Choose a location for the file and click the SAVE button.

When exporting a comma delimited report, add .csv to the end of the filename to easily open the report in Excel.

10. The amount of time required to create the report varies depending on the number of samples and items selected. An “Export Complete” message displays when the report is finished. Click the OK button.

ACCESSING THE SAMPLE ENTRY SCREEN

Use the SAMPLE ENTRY Screen to create new sample records or update existing sample

records.

Access the SAMPLE ENTRY Screen by clicking the Samples Menu and then clicking the

Sample Entry command.

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Figure 16: The SAMPLE ENTRY Screen.

The “Business Unit List of Values” pop-up window displays for users authorized for more than one business unit. Select the relevant Business Unit to enter samples for and then click the OK button.

CREATING SAMPLE RECORDS

This section provides steps for creating sample records in SIS using the SAMPLE ENTRY

Screen. The Sample Header and Sampling Conditions TAB contain the only required fields.

The rest of the fields are optional and should be entered when applicable.

Sample Header

The sample header is required. It identifies the general sample information including collector,

date and time collected, sequence number, and sampling reason.

Figure 17: A sample header.

1. Access the SAMPLE ENTRY Screen. 2. Enter the four-digit collector number assigned to the employee, group, or monitoring

device that collected the sample in the Collector ID field. Press the [TAB] key. 3. Enter the date the sample was collected in the first Date Collected field (format

MM/DD/YYYY). Press the [TAB] key. 4. Optionally, enter the time the sample was collected in military time (i.e., enter 1:00 pm

as 1300, enter 9:00 am as 0900) in the second Date Collected field. Press the [TAB] key.

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5. Enter the sequence number for the sample in the Sequence# field. Press the [TAB] key.

6. Click the list of values button next to the Reason field to select a reason for sampling. The field defaults to “Routine Sampling.”

Project/Facility/Monitoring Point TAB

This section is used to indicate if the collected sample is related to a project, eFACTS facility,

or a monitoring point.

1. Click the Project/Facility/Monitoring Pt TAB.

Figure 18: The Project/Facility/Monitoring Pt TAB.

2. If the sample was collected for an existing project, complete the following steps: a. Enter the code identifying the project’s business unit or select using the list of

values button to the right of the Project Unit field. Press the [TAB] key. b. Enter the identification number assigned to the project or select using the list of

values button to the right of the Project ID field. 3. If the sample was collected to monitor an existing primary facility and/or sub facility,

complete the following steps: a. Enter the program-specific identification number in the Primary Facility field or

select a primary facility from the list of values by entering a facility name or program. Press the [TAB] key.

b. Optionally, select a sub-facility by clicking the list of values button to the right of the Sub-Facility field.

4. If the sample was collected at a particular monitoring point, identify the monitoring point using one of the following methods:

a. Enter the Monitoring Point ID assigned to the monitoring point and press the [TAB] key.

b. Enter the Monitoring Point Alias and press the [TAB] key. c. Click the list of values button to the right of either Monitoring Point field, enter a

latitude and longitude, and click the ACCEPT button to display a list of monitoring points to select from.

Coordinates are entered in the format DD-MM-SS.SSSS (e.g., 41-29-46.3034 or -79-38-09.6071)

5. If a monitoring point was entered, click the SAVE button on the toolbar or press the [F10] key. If necessary, proceed to the next tab.

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6. If a monitoring point was not entered, proceed to the Sampling Conditions TAB.

Sampling Conditions TAB

This section is used to enter the conditions under which the sample was collected. The

Medium Type and Medium fields are required. If the sample is linked to a monitoring point on

the Project/Facility/Monitoring Pt TAB, the Medium Type and Medium fields are populated

using information from the monitoring point.

1. Click the Sampling Conditions TAB.

Figure 19: The Sampling Conditions TAB.

2. The Medium Type field is required. Enter the code that identifies the type (category) of sample medium (soil, water, air, plants, etc.) or select by using the list of values button to the right of the Medium Type field. Press the [TAB] key.

3. The Medium field is required. Enter the code that identifies the sample medium or select by using the list of values button to the right of the Medium field. Press the [TAB] key.

4. The fields that display vary based on the Business Area selected when first accessing the SAMPLE ENTRY Screen. Enter data in the optional fields as necessary.

5. Click the SAVE button on the toolbar or press the [F10] key. 6. If necessary, proceed to the next TAB.

Location TAB

This section is used to identify the location at which a sample was collected.

The latitude, longitude and datum are required in order to link an NHD record to a sample.

If a sample is linked to a monitoring point on the Project/Facility/Monitoring Pt TAB, the

Locational TAB automatically displays the locational information from the monitoring point.

B-27

Figure 20: The Location TAB.

1. Click the Location TAB. 2. Click the AUTO-FILL button (The county and municipality displays based on the linked

primary facility, sub facility, or monitoring point). If the county and municipality does not display, complete Steps c through e; otherwise, proceed to Step f.

3. The state defaults to “PA”. Update if necessary. Press the [TAB] key. 4. Select the county by using the list of values button. 5. Select the municipality by using list of values button. 6. Select the quadrangle by using the list of values button. 7. (Required to insert NHD) Enter the latitude where the sample was taken in the Latitude

field. Press the [TAB] key. 8. (Required to insert NHD) Enter the longitude where the sample was taken in the

Longitude field. Press the [TAB] key.


9. Enter the UTM Zone where the sample was taken in the UTM field. Press the [TAB] key.

10. Enter the northing where the sample was taken in the Northing field. Press the [TAB] key.

11. Enter the easting where the sample was taken in the Easting field. Press the [TAB] key. 12. (Required to insert NHD) Enter the horizontal reference datum used to calculate the

point at which the sample was collected or select by using the list of values button to the right of the Datum field. Press the [TAB] key.

13. Enter the method used to identify the point at which the sample was collected or select by using the list of values button to the right of the Location Method field. Press the [TAB] key.

14. Enter a description of the location at which the sample was collected in the Location field.

15. Click the SAVE button on the toolbar or press the [F10] key. 16. Proceed with linking the sample to an NHD record or to the next tab.

Linking the Sample to a NHD Record

This section identifies the steps for linking a sample to a NHD record. Complete the step for

entering details on the Location TAB before linking the sample to a NHD record.

B-28

If the sample is associated with a monitoring point, the NHD record for the monitoring point will “automatically” display for the sample and cannot be updated. Therefore, this procedure cannot be completed.

1. Click the Location TAB. 2. Verify that the latitude, longitude, and datum exist for the sample. If not, insert the

latitude, longitude, and datum.

To insert an NHD record for a sample, the latitude, longitude and datum must be entered on the Location TAB.

3. Click the GET/VIEW NHD DATA button at the bottom of the Location TAB. The NHD pop-up window displays.

Figure 21: The NHD pop-up window.

4. Click the LAUNCH NHD LOCATOR button at the bottom of the screen.

Figure 22: The NHD locator.

5. Use the NHD Locator Tool to either accept the default snapped point or create a user-defined, new snapped point to accept. Reference the NHD Locator Tool User Guide for the steps.

B-29

6. Click the ACCEPT SNAPPED POINT(S) button and then click the OK button to exit the NHD Locator Tool and return to the SAMPLE ENTRY Screen.

7. Click the GET NHD DATA button to add the NHD record created via the NHD Locator Tool to the sample.

8. Click the CLOSE button to return to the Location TAB.

Name and Address TAB

This section is used to enter a name and address associated with the collection location of the

sample.

1. Click the Name & Address TAB.

Figure 23: The Name & Address TAB.

2. Enter the name of the location where the sample was collected in the Addressee field. Press the [TAB] key.

3. Enter the first line of the address where the sample was collected on the first line in the Address field. Press the [TAB] key.

4. If necessary, enter the second line of the address where the sample was collected in the Address field. Press the [TAB] key.

5. Enter the city where the sample was collected in the City field. Press the [TAB] key. The state defaults to “PA”.

6. Enter the zip code where the sample was collected in the Zip Code field. Press the [TAB] key.

7. Enter the phone number where the sample was collected in the Phone# field. 8. Click the SAVE button on the toolbar or press the [F10] key. 9. If necessary, proceed to the next tab.

Comment/Quality Assurance TAB

This section is used to enter the comments and quality assurance details regarding the

sample.

1. Click the Comments/Quality Assurance TAB.

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Figure 24: The Comments/Quality Assurance TAB.

2. Enter a description of the sample’s appearance. Press the [TAB] key. 3. Enter any additional information regarding the sample. Press the [TAB] key. 4. Enter the quality assurance type (duplicate, blank, or spike). 5. If a duplicate, click the list of values button, enter the ID of the collector for the duplicate

sample, the date the duplicate was collected, and then click the ACCEPT button. 6. If necessary, select a confidentiality reason code (e.g., private water supply or legal

enforcement action). 7. If the sample is to be voided due to quality assurance reasons, select the “Void Sample”

checkbox. 8. If the sample is to be dry, select the “Dry Sample” checkbox. 9. Click the SAVE button on the toolbar or press the [F10] key. 10. If necessary, proceed to the next tab.

Field Tests TAB

This section is used to identify the types of tests conducted in the field on the sample.

1. Click the Field Tests TAB.

Figure 25: The Field Tests TAB.

2. Use the scrollbar to locate the field test for the results and click in the Result Amount field.

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The list of field tests varies based on the Business Unit.

3. Enter the amount for the test. Press the [TAB] key. 4. Update the unit of measurement if necessary. 5. Repeat the previous steps all applicable field tests are entered. 6. Click the SAVE button on the toolbar or press the [F10] key.

Inserting Additional Samples for the Same Location

This section identifies how to enter an additional sample at the same location as the previously

entered sample.

1. While the previous sample is displayed on the screen, click the SAME LOCATION button. A new record matching the header, project, facility, monitoring point, and location details of the previous sample is created.

2. Update the sequence number. 3. Update the sample detail tabs as described in the CREATING SAMPLE RECORDS

section of the user guide.

Inserting Additional Samples at a New Location

To enter an additional sample at a new location:

1. While the previous sample is displayed on the screen, click the NEW LOCATION button. A new record with header details matching the previous sample will be created.

2. Update the sequence number and reason. 3. Update the sample detail tabs as described in the CREATING SAMPLE RECORDS

section of the user guide.

Voiding a Sample

1. Access the SAMPLES Screen by clicking the Samples Menu and then clicking the Query sample command.

2. Query the sample to be voided. 3. Click the Comments/Quality Assurance TAB.

Figure 26: The Comments/Quality Assurance TAB.

4. If applicable, select a quality assurance type to identify a reason for voiding the sample. Press the [TAB] key.

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If the quality assurance type is duplicate and only one sample record exists for the collector and date collected, the duplicate sample automatically displays. Otherwise, click the list of values button to identify the duplicate.

5. If applicable, select the Void Sample checkbox. 6. If applicable, select the Dry Sample checkbox. 7. Click the SAVE button on the toolbar or press the [F10] key.

UPDATING SAMPLE RECORDS

Update existing sample records using the SAMPLE ENTRY Screen.

1. Access the SAMPLE ENTRY Screen by clicking the Samples Menu and then clicking the Sample Entry command.

2. Click the ENTER QUERY button on the toolbar and query for a sample as described in the

3. QUERYING FOR SAMPLES section of the user guide.

4. Update the fields described in the CREATING SAMPLE RECORDS section of the user guide.

5. When finished click the SAVE button on the toolbar or press the [F10] key.

MONITORING POINTS

This section provides steps for how to create a new monitoring point using the MONITORING

POINTS Screen. Access the MONITORING POINTS Screen by clicking the Samples Menu

and then clicking the Monitoring points command.

Use monitoring points to indicate a consistent sampling location. Monitoring points can also be

linked to an eFACTS facility or a SIS project.

Figure 27: The MONITORING POINTS Screen.

Creating a New Monitoring Point

Step 1: Header Block (required)

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Use the header block to identify the monitoring point name and type.

Figure 28: The monitoring point header block.

1. Click the CREATE RECORD button on the toolbar or press the [F6] key. 2. Enter the name of the monitoring point and press the [TAB] key. 3. Enter the code that identifies the type of monitoring point or select one using the list

of values button. Step 2: Location TAB (required)

Use the Location TAB to identify the location of the monitoring point from a general location

description to specific coordinates. The following fields are required:

• State

• County

• Latitude

• Longitude

The Datum field is required if linking an NHD record to the monitoring point.

1. Click the Location TAB.

Figure 29: The Location TAB.

2. The state defaults to “PA”. Update if necessary. 3. Select a code identifying the county in which the sample was collected using the list

of values button. 4. Optionally, select the code identifying the municipality in which the sample was

collected by using the list of values button. 5. Enter the latitude where the sample was taken in Degrees-Minutes-Seconds (e.g.,

40-15-42.8394). 6. Enter the longitude where the sample was taken in Degrees-Minutes-Seconds (e.g.,

-76-52-46.9194). 7. Optionally, enter the horizontal reference datum used to calculate the point at which

the sample was collected or select by using the list of values button. 8. Complete any of the optional fields with available information:

• Municipality

B-34

• Quadrangle

• Datum

• Reference Point

• Geometric Type

• Location

• Depth

• Elevation

• Accuracy

• Altitude Datum

• UTM Zone

• Northing

• Easting

Click in a field to display a description in the hint line.

Step 3: Function/Administration TAB (required)

Use the Function/Administration TAB to identify the function, medium type, and sample

medium for a monitoring point as well as administrative details such as the closed date for a

monitoring point. The Medium Type and Sample Medium fields are required.

1. Click the Function/Administration TAB.

Figure 30: The Function/Administration TAB.

2. Optionally, enter a code identifying the function of the monitoring point or select by using the list of values button.

3. Enter the code identifying the sample medium type monitored by the monitoring point or select by using the list of values button.

4. Enter the code identifying the sample medium monitored by the monitoring point or select by using the list of values button.

5. If the monitoring point is a STORET station, select the STORET Station checkbox. 6. If the monitored material is treated, select the Monitored Material Treated checkbox. 7. If monitoring gages are associated with the monitoring point, complete the following

steps: a. Click the GAGES button and the “Monitoring Gages” pop-up window

displays.

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Figure 31: The “Monitoring Gages” pop-up window.

b. To go to a blank record, click the CREATE RECORD button on the toolbar or press the [F6] key.

c. Enter the gage location ID or select by using the list of values button. d. Repeat Steps ii and iii until all associated gages are selected. e. Click the CLOSE button.

8. Enter a code identifying the source of the information regarding the monitoring point (e.g., home owner, DEP, etc.) or select by using the list of values button.

9. Enter a code identifying the method used to identify the point coordinates of the monitoring point or select by using the list of values button.

10. Enter the map scale used to locate the monitoring point. 11. Enter the accuracy measurement of the coordinates and select a code identifying

the unit of measure for the accuracy measurement using the list of values. 12. Click the SAVE button on the toolbar or press the [F10] key.

Step 4: Aliases TAB (optional)

Use the Aliases TAB to create identifiers for the monitoring point in addition to the ID

generated by SIS.

1. Click the Aliases TAB.

Figure 32: The Aliases TAB.

2. Click the CREATE RECORD button on the toolbar or press the [F6] key to add a new record.

3. Enter the alias ID for the monitoring point. Press the [TAB] key. 4. Enter a description of the alias. 5. Repeat Steps b through d to enter additional aliases.

Linking the Monitoring Point to a NHD Record

Link a monitoring point to a NHD record to monitoring the environmental impact on a water

resource.

1. Query an existing monitoring point or create a new one.

B-36

2. Click the Location TAB. 3. Verify that the latitude, longitude, and datum are entered. 4. Click the NHD TAB.

Figure 33: The NHD TAB.

5. Click the LAUNCH NHD LOCATOR button at the bottom of the screen and the NHD Locator Screen displays.

Figure 34: The NHD locator.

6. Use the NHD Locator Tool to either accept the default snapped point or create a user-defined, new snapped point to accept.

7. Click the ACCEPT SNAPPED POINT(S) button and then click the OK button to exit the NHD Locator Tool and return to the SAMPLE ENTRY Screen.

8. Click the GET NHD DATA button to add the NHD record created via the NHD Locator Tool to the sample.

9. Click the SAVE button on the toolbar or press the [F10] key.

B-37

Linking Primary Facilities to a Monitoring Point

Link a monitoring point to an eFACTS primary facility to monitor its environmental impact on

the soil, water, or air.

Caution: Do NOT link the monitoring point to a primary facility if intending to link to specific

sub-facilities of that primary facility. Instead proceed to the section on linking sub-facilities.

1. Query an existing monitoring point or create a new one. 2. Click the PRIMARY FACILITIES button and the “Monitoring Point/Primary Facilities” pop-up

window displays. 3. Click the CREATE RECORD button on the toolbar or press the [F6] key.

Figure 35: The “Monitoring Point/Primary Facilities” pop-up window.

4. Select a primary facility by entering the “Other ID” from eFACTS (i.e., permit, registration, or license number) or search for a primary facility by name. To search for a primary facility by name:

a. Click the list of values button to display the “List Reduction Criteria” pop-up window.

b. Enter the name of the facility (use the wildcard symbol if necessary) c. Optionally, select a program to limit the list of matching primary facilities. d. Click the ACCEPT button and the “FIX Primary Facilities” pop-up window

displays. e. Select the correct facility from the list and click the OK button.

5. Optionally, select an Alias ID to associate this primary facility with one of the aliases created for the monitoring point

6. Update the effective date if necessary. 7. Click the SAVE button on the toolbar or press the [F10] key. 8. Click the CLOSE button.

Linking Sub-facilities to a Monitoring Point

Link a monitoring point to an eFACTS sub-facility to monitor its environmental impact on the

soil, water, or air.

1. Query an existing monitoring point or create a new one. 2. Click the SUB-FACILITIES button and the “Monitoring Points/Sub-Facilities” pop-up window

displays. 3. Click the CREATE RECORD button on the toolbar or press the [F6] key.

B-38

Figure 36: The “Monitoring Point/Sub-Facilities” pop-up window.

4. Select a primary facility by entering the “Other ID” from eFACTS (e.g., permit, registration, license) or search for a primary facility by name.

To search for a primary facility by name:

a. Click the list of values button to display the “List Reduction Criteria” pop-up window. b. Enter the name of the facility (use the wildcard symbol if necessary) c. Optionally, select a program to limit the list of matching primary facilities. d. Click the ACCEPT button and the “FIX Primary Facilities” pop-up window displays. e. Select the correct facility from the list and click the OK button.

5. Optionally, select an Alias ID to associate this primary facility with one of the aliases created for the monitoring point

6. Select a sub-facility using the list of values button. 7. Update the effective date if necessary. 8. Click the SAVE button on the toolbar or press the [F10] key. 9. Click the CLOSE button.

Linking Projects to a Monitoring Point

Link a monitoring point to a project to monitor its environmental impact on the soil, water, or

air.

1. Query an existing monitoring point or create a new one. 2. Click the PROJECTS button and the “Monitoring Point/Projects” pop-up window displays.

Figure 37: The “Monitoring Point/Projects” pop-up window.

3. Click the CREATE RECORD button on the toolbar or press the [F6] key. 4. Optionally, enter a business unit code in the Project Unit field or select one using the list of

values. 5. Enter a Project ID or select one using the list of values. 6. Optionally, select an Alias ID to associate this project with one of the aliases created for the

monitoring point 7. Update the effective date if necessary. 8. Click the SAVE button on the toolbar or press the [F10] key.

Closing a Monitoring Point

Closing monitoring point disallows linking samples to it.

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1. Query an existing monitoring point. 2. Click the Function/Administration TAB.

Figure 38: The Function/Administration TAB.

3. Enter a date in the Date Closed field. Press the [TAB] key. 4. Enter a comment regarding the closure. 5. Click the SAVE button on the toolbar or press the [F10] key.

PROJECTS

Use projects to relate samples collected for a specific purpose or to group samples or

monitoring points together for the purposes of query sample results or running reports. Link a

sample to a project using the SAMPLE ENTRY Screen or link a monitoring point to a project

using the MONITORING POINTS Screen.

Querying for Existing Projects

1. Click the Samples Menu and then click the Projects command. The PROJECTS Screen displays.

2. Click the ENTER QUERY button on the toolbar or press the [F7] key. 3. Enter exact text criteria or use the wildcard symbol (%) in the Project ID, Description,

and/or Comments fields or select a business unit or cost center using the lists of values.

The Comments field is case-sensitive. The Project ID and Description fields are not.

4. Click the COUNT HITS button or press [SHIFT] + [F2] to display the number of matching projects in the hint line.

5. Click the EXECUTE QUERY button or press the [F8] key to display the matching projects.

Inserting a New Project

1. Click the Samples Menu and then click the Projects command. The PROJECTS Screen displays.

2. Enter a unique project ID and press the [TAB] key. 3. Enter a project description and press the [TAB] key. 4. Enter a business unit or select one using the list of values. Press the [TAB] key. 5. Enter a cost center or select one using the list of values. 6. Optionally, enter any comments about the purpose of the project. 7. Click the SAVE button on the toolbar or press the [F10] key.

B-40

REPORTS

Requesting Sample Reports

After lab analysis of a sample is complete, a report is e-mailed to the collector, business unit,

and report groups linked to the sample. Reports for groups of samples collected during a date

range or identified by collector ID and sequence numbers can be e-mailed again or sent to a

different address by requesting sample reports.

1. Click the Reports Menu, select Sample reports, and then click the Request sample reports command. The REQUEST SAMPLE REPORTS Screen displays.

2. The screen is automatically placed into query mode. Enter a collector ID, collection date, and/or sequence number to locate a particular set of samples.

To enter a range of collection dates or sequences, click the QUERY OPTIONS button instead of specifying a specific date or sequence.

3. Click the COUNT HITS button on the toolbar to display the number of samples matching the query criteria in the hint line.

4. After verifying the expected or appropriate number of samples will be retrieved, click the EXECUTE QUERY button on the toolbar to display the matching samples.

5. Select the samples to have reports generated by clicking the “Select Sample” checkbox or click the SELECT ALL button.

6. Select either the default report destination or a custom report destination.

Click the SHOW DEFAULT button to see the default report destination. Click the CUSTOMIZE button to enter a custom report destination. The custom report destination defaults to the current user.

7. Click the PRINT REPORTS button and a confirmation dialog displays indicating the selected reports have been e-mailed to the destination specified. Click the OK button.

Generating Sample Reports

The sample reports for samples collected by a specific collector during a date range can be

generated and viewed in a PDF file.

1. Click the Reports Menu, select Sample reports, and then click the Long/Short Sample report command. The LONG/SHORT REPORTS Screen displays.

2. The short report type is selected by default. Select the desired report type.

The short sample report form displays on a page with a portrait orientation and has a compact layout. The long sample report displays on a page with a landscape orientation and typically uses more pages for each sample than the short report.

3. Enter a collector ID. 4. Enter a beginning sample collection date in the SAMPLE BEGIN DATE field. 5. If necessary, change the numbers in the BEGIN and END SEQUENCE NO. fields 6. Click the RUN REPORT button. After a period of time that varies depending on the number

matching sample reports, a PDF will open with the sample reports.

B-41

Missing Sample Header/Results Report

Use the Missing Sample Header/Results report to view a list of sample records entered into

SIS that do not have a lab results or to view a list of lab results available in SIS that have not

had a sample record entered.

1. Click the Reports Menu and then click the Missing sample header/results command. The MISSING HEADER/RESULTS REPORTS Screen displays.

2. Select the Missing Header report to generate a list of sample reports awaiting entry of the sample in SIS or select the Missing Result report to generate a list of samples in SIS awaiting sample reports.

3. Select a Collector Group, Business Unit, or Collector to run the report for from the lists of values.

4. Enter a collection date range for either the Missing Header or Missing Results report. 5. Click the RUN REPORT(S) button.

Monitoring Points Report

The Monitoring Points report lists all of the monitoring points linked to a specified project,

primary facility, or sub-facility including the location of the monitoring point, whether it is active,

and the date last sampled.

1. Click the Reports Menu and then click the mOnitoring points command. The MONITORING POINT REPORTS Screen displays.

2. Select either a PDF or Excel file for the report destination. 3. Select report type to indicate whether the monitoring points will be identified by project,

primary facility, or sub-facility. If running a report by project:

a. Optionally, identify a project unit to narrow down the list of projects. b. Select a project ID. c. Click the RUN REPORT button.

If running a report by primary facility:

a. Enter the eFACTS “Other ID” for a primary facility. The “Other ID” is typically a permit or registration number assigned by the related program.

b. Alternatively, use the list of values to select a facility by either facility name or associated program.

c. Click the RUN REPORT button. If running a report by sub-facility:

a. First, enter the eFACTS “Other ID” for a primary facility. The “Other ID” is typically a permit or registration number assigned by the related program.

b. Alternatively, use the list of values to select a primary facility by either facility name or associated program.

c. Select the sub-facility using the list of values. d. Click the RUN REPORT button.

4. Optionally, select the checkbox to include the date each monitoring point was last sampled.

Including the date last sampled for each monitoring point lists the most recent sample regardless of which business unit collected the sample. Use the list of values to restrict it to a specific business unit.

Module 8.1A Report

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SIS can generate Module 8.1A reports for a specific monitoring point related to a project or

eFACTS primary facility or for all monitoring points between a pair of latitudes and longitudes.

To run the report for an individual monitoring point related to a project or eFACTS primary

facility:

1. Click the Reports Menu and then click the module 8.1A command. The MODULE 8.1A REPORTS Screen displays.

2. Select either a PDF or Excel file for the report destination. 3. Select a report type to indicate whether the monitoring points will be identified by project or

primary facility. If running a report by project:

a. Optionally, identify a project unit to narrow down the list of projects. b. Select a project ID. c. Select a monitoring point alias from the MP Alias list of values.

If running a report by primary facility:

a. Enter the eFACTS “Other ID” for a primary facility. The “Other ID” is typically a permit or registration number assigned by the related program.

b. Alternatively, use the list of values to select a facility by either facility name or associated program.

c. Select a monitoring point alias from the MP Alias list of values. 4. Enter a beginning and ending sample collection date range. 5. Optionally, identify to include only samples collected by a specific business unit. 6. Click the RUN REPORT button.

To run the report for all monitoring points between a pair of latitudes and a pair of longitudes:

1. Click the Reports Menu and then click the module 8.1A command. The MODULE 8.1A REPORTS Screen displays.

2. Select either a PDF or Excel file for the report destination. 3. Select the By Latitude/Longitude report type. 4. Enter the latitude values in the low and high fields. 5. Enter the longitude values in the low and high fields.


6. Enter a beginning and ending sample collection date range. 7. Optionally, identify to include only samples collected by a specific business unit. 8. Click the RUN REPORT button.

B-43

B.4 SEDIMENT FIELD DATA COLLECTION SHEET

B-46

B.5 CIM DEPLOYMENT FORM

B-49

B.6 CIM MAINTENANCE VISIT FIELD FORM

B-54

B.7 TEMPERATURE ADJUSTMENTS FOR PH BUFFERS

B-55

Table 1. Temperature adjusted pH buffer values from British Drug House (BDH;

“Temperature Dependence of pH,” n.d.)

TEMP (C) 0 5 10 15 20 25 30 35 40 45 50

BDH5022 4.00 4.00 4.00 4.00 4.00 4.00 4.01 4.02 4.03 4.04 4.06

BDH5050 7.12 7.09 7.06 7.04 7.02 7.00 6.99 6.98 6.98 6.97 6.97

BDH5076 10.31 10.23 10.17 10.11 10.05 10.00 9.95 9.91 9.87 9.81

4 Buffer y = 0.0000357143x2 - 0.0003928571x + 3.9885714286

7 Buffer y = 0.0000680653x2 - 0.0063487179x + 7.1191608392

10 Buffer y = 0.0000954898x2 - 0.0147159972x + 10.3068957012

Figure 1. Second-order polynomial regression lines fitted to temperature-corrected pH

values for BDH buffers.

3.96

4.00

4.04

4.08

0 10 20 30 40 50 60

pH

Va

lue

Temperature (C)

6.95

7.00

7.05

7.10

7.15

0 10 20 30 40 50 60

pH

Va

lue

Temperature (C)

9.60

9.80

10.00

10.20

10.40

0 10 20 30 40 50 60

pH

Va

lue

Temperature (C)

B-57

B.8 DO 100% SATURATION CHART FOR FRESH WATER

C-1

APPENDIX C: PHYSICAL DATA COLLECTION INFORMATION AND FORMS

C-2

C.1 RIFFLE/RUN PREVALENCE HABITAT DATA COLLECTION FORM

C-5

C.2 RIFFLE/RUN PREVALENCE HABITAT DATA COLLECTION ABBREVIATED (ONE-PAGE)

FORM

C-7

C.3 LOW GRADIENT HABITAT DATA COLLECTION FORM

C-10

C.4 PEBBLE COUNT DATA COLLECTION FORM

C-11

PEBBLE COUNT FORM GIS Key:

Survey Crew:

Stream:

County:

SWP:

Mean Width:

Sample Interval:

Reach Length:

Station Description:

1 35 69 102 135 168

2 36 70 103 136 169

3 37 71 104 137 170

4 38 72 105 138 171

5 39 73 106 139 172

6 40 74 107 140 173

7 41 75 108 141 174

8 42 76 109 142 175

9 43 77 110 143 176

10 44 78 111 144 177

11 45 79 112 145 178

12 46 80 113 146 179

13 47 81 114 147 180

14 48 82 115 148 181

15 49 83 116 149 182

16 50 84 117 150 183

17 51 85 118 151 184

18 52 86 119 152 185

19 53 87 120 153 186

20 54 88 121 154 187

21 55 89 122 155 188

22 56 90 123 156 189

23 57 91 124 157 190

24 58 92 125 158 191

25 59 93 126 159 192

26 60 94 127 160 193

27 61 95 128 161 194

28 62 96 129 162 195

29 63 97 130 163 196

30 64 98 131 164 197

31 65 99 132 165 198

32 66 100 133 166 199

33 67 101 134 167 200

34 68

Comments:

C-12

C.5 ALTERNATIVE PEBBLE COUNT DATA COLLECTION FORM

D-1

APPENDIX D: PENNSYLVANIA’S SURFACE WATER QULAITY MONITORING NETWORK

(WQN) MANUAL

D-3

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D-4



PENNSYLVANIA ’S SURFACE WATER QUALITY MONITORING NETWORK (WQN) MANUAL

2020

D-5

Originally prepared: 1972

Update prepared by:

Molly Pulket






Edited by:

Molly Pulket and Josh Lookenbill






2019

Molly Pulket and Josh Lookenbill






2020

D-6

INTRODUCTION

The Pennsylvania Water Quality Network (WQN) is a statewide, fixed-station water quality sampling

system operated by the Pennsylvania Department of Environmental Protection’s (DEP) Bureau of

Clean Water (BCW). It is designed to track both the quality of Pennsylvania’s surface waters and the

effectiveness of the water quality management program by accomplishing four basic objectives:

• Monitor temporal water quality trends in major surface streams throughout Pennsylvania.

• Monitor temporal water quality trends in selected reference waters.

• Monitor the trends of nutrient and sediment loads in the major tributaries entering the

Chesapeake Bay.

• Monitor temporal water quality trends in selected Pennsylvania lakes.

“Major” streams, for the purposes of this document, are interstate and intrastate waters with drainage

areas of roughly 200 square miles or greater. These waters receive both point and non-point source

pollutants and are sampled at or near their mouths to measure overall quality before flows enter the

next higher order stream or before exiting the state. In this way, trends can be established, and the

effectiveness of water quality management programs can be assessed by watershed. For more

detailed information on the objectives of the WQN, see Subappendix D.5.

The monitoring of the WQN is conducted collaboratively between DEP, USGS Pennsylvania Water

Science Centers, and Susquehanna River Basin Commission (SRBC).

WQN resources can be found on DEP’s WQN webpage: www.dep.pa.gov, keywords: Water Quality

Network.

STATION TYPES

The WQN consists of 177 active stations grouped into three primary station types: standard,

Chesapeake Bay nutrient loading, and reference.

The 116 standard stations are generally sampled every other month for stream discharge

measurements and chemical analysis, and every other year for a biological and physical evaluation. Of

these, 28 are sampled monthly for chemical analysis and 17 are considered low alkalinity stations

which are analyzed for a few additional parameters. See Subappendix D.1 for the list of parameters

collected at low alkalinity stations.

The 36 Chesapeake Bay Nutrient Loading (Chesapeake Bay) stations are sampled monthly for stream

discharge measurements and chemical analysis, and once a year for biological and physical

evaluation. These stations are also part of the Chesapeake Bay Program nontidal monitoring network

(CBP NTN) which requires eight additional water chemistry samples to be collected annually during

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storm events. More information on the CBP NTN can be found at:

CBP nontidal water quality monitoring.

The 23 reference stations are sampled monthly for stream discharge measurements and chemical

analysis and typically once a year (spring or fall) for biological and physical evaluation. For method

development purposes, some reference stations are sampled twice a year, once in the spring and once

in the fall. Reference stations are selected to represent minimally disturbed waters within

Pennsylvania’s various bioregions. Bioregions are areas with similar water quality characteristics

because of like features such as topography, land use, soils, and potential natural vegetation. Although

it may be argued that Pennsylvania has no unaffected waters (by atmospheric deposition), reference

stations reasonably describe water quality that could be achieved in the absence of point source

discharges and in the presence of minimal or typical non-point source impacts. Thus, these stations

can provide, in the absence of site-specific data, a benchmark for use in the water quality management

permitting program as well as a measure of changing water quality in areas of the state not directly

impacted by development. Of the 23 reference stations, 16 are monitored for atmospheric deposition

and therefore, are analyzed for a few additional parameters (Subappendix D.1).

WQN lake stations (excludes Lake Erie and Presque Isle Bay which are standard WQN stations) are

selected after consideration of size, public access, intensity of use, and availability of existing data.

Large lakes with heavy public use and/or historical data are favored for inclusion in this program

because changing trends in the water quality of these resources have the potential for serious impacts

on water uses. Approximately 90 lakes are included in this network, however, sampling occurs in

groups of 15-20 lakes on a rotation. These lake groups are sampled once a year for five consecutive

years before initiating a new group. The five-year data sets are then used to assess lake water quality

trends. Monitoring at the current group of 15 lakes started in 2016 and will continue through 2020.

Station specific details are included in the Active, Macroinvertebrate, and Discontinued Stations tables

found in Subappendices D.2-D.4. A web mapping application is also available to view each station at

WQN web application.

UPDATES TO THE WQN

This revision of the WQN manual covers changes that occurred from October 2018 through September

2020. All information in this edition is reflective of the WQN’s status as of October 1st, 2020.

Dissolved arsenic, nitrate, and nitrite were removed from Standard Analysis Code (SAC) 015. While six

new parameters were added to SAC 015: Ammonia D, NO2 + NO3 T, NO2 + NO3 D, Phosphorus D,

Ortho Phos D, and DOC. All SAC 013 stations will use the new, modified SAC 015 to simplify data

collection. A new SAC 122 was created to provide additional information associated with atmospheric

deposition effects at 16 of the reference stations. All lake stations that used SAC 017 will now use SAC

083. All SACs used for the WQN and the parameters they contain are listed in Subappendix D.1. The

SACs assigned to each WQN station can be found in Subappendices D.2 and D.4 (Active and

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Discontinued Station lists). In September 2020, the five-year rotation of the reference stations ended,

and 23 stations were selected for the Water Year (WY) 2021-2025 rotation. Of these 23, 14 reference

stations were retained to build datasets that span more than five years, eight new stations were added,

and one station was reactivated. Two new standard stations were created on Lick Run to aid in

limestone protocol and method development and two standard stations were reactivated to aid in

permitting decisions and part of the effort to collect Perfluoroalkyl and polyfluoroalkyl substances

(PFAS) data. Standard stations 467, 635, and 843 were discontinued. The WQN specific sample

submission form will be phased out and the standard DEP Bureau of Laboratories (BOL) submission

forms will be used.

The tables and subappendices in this update reflect these changes.

WATER CHEMISTRY SAMPLING

Water chemistry samples are collected at all active WQN stations based on the frequency assigned in

the Active Stations table (Subappendix D.2). Reference and Chesapeake Bay stations are sampled

once every month. The majority of standard stations are sampled once every other month and lakes

are sampled once annually. Sampling for a station, no matter the station type, should be on a fixed

scheduled. This allows for the potential of a wide range of sampling conditions.

At least one Standard Analysis Code (SAC) is assigned to each WQN station depending on the station

type. A SAC contains a list of analytes and is used to identify which water chemistry tests should be

performed on a sample. A list of the SACs and the station type associated with it can be found in

Subappendix D.1. Selected water chemistry tests may be added to any of the WQN stations on a case-

by-case basis when the need for additional data is determined. However, the use of additional analyses

for extended periods of time is discouraged in favor of creating a new SAC tailored to the needs of the

collector. All such changes are coordinated with the DEP BOL through the Bureau of Clean Water’s

Water Quality Division (WQD).

Chemical Sampling Procedures

Multiple agencies and numerous field staff are part of the WQN sampling effort. It is important that all

involved are using the same data collection protocols to ensure reliable and comparable data. Any

questions regarding sampling protocols should be directed to the DEP WQD, 717.787.9637.

All WQN collectors must obtain a four-digit collector identification number assigned by BOL. The

Collector ID Request Form (ID Request Form) should be filled out and submitted via email to the WQD.

The “DEP Laboratory Submission Sheet” (Submission Sheet) must be filled out and submitted to the

BOL. The information required on this form varies between station types. Field staff are responsible for

working with the WQD to ensure the form is filled out correctly.

D-9

Surface water samples are collected as isokinetic, depth integrated, single mid-channel or equal-width

samples. Samples are normally collected in High-Density Polyethylene (HDPE) or amber glass (for

organic compound analysis) sample bottles. A sample volume and field preservation chart are available

for reference in Subappendix D.6. Bottles must be labeled with the WQN number, collector ID number,

sequence number, date, and time. If space allows, adding the SAC, general test descriptions (total

metals, TOC, etc.), filtered, and preservation type can help minimize any questions concerning the

sample analyses. Depending on the station, samples may be collected from a bridge, boat, or by

wading.

Detailed descriptions on how to collect surface water samples can be found in Chapter 4 of DEP’s

Water Quality Monitoring Protocols for Streams and Rivers (the Monitoring Book, Shull and Lookenbill

2021) and in Chapters 4 and 5 of the USGS’s National Field Manual for the Collection of Water Quality

Data (USGS 2006, Wilde 2004).

Standard Stations

A standard station is typically assigned SAC 010 or 610 and can be located anywhere in Pennsylvania.

SACs 016 and 616 are used if the watershed experiences low alkalinity. The 610 and 616 SACs are

used for stations in geographic areas that could be underlain by Marcellus shale; these SACs contain a

few additional analytes typically associated with Marcellus formation water. Reason Code 01 should be

used on the Sample Submission Sheet for all standard stations.

Reference Stations

These stations are assigned either SAC 015 or the Atmospheric Deposition SAC 122, depending on

drainage area and location. Both SACs contain the analyte dissolved organic carbon (DOC). The

analytical method for DOC requires that a blank sample is associated with every DOC sample. To

satisfy this requirement, a blank DOC sample is collected with the first SAC 015 or 122 sample in a

sampling group. A sampling group can contain the samples collected in the same day, week, project,

etc. but should not be more than 20 samples. The collector ID and sequence number for that blank

should be recorded on all the Sample Submission forms for that sampling group and specify that it is

the blank to be associated with these DOC samples. A new blank DOC sample should be collected

with each sampling group. Reference stations are additionally sampled for fecal coliforms and E. coli; a

sterilized 125 ml bacteriological bottle pretreated with sodium thiosulfate (neutralizes the effects of

residual chlorine) is used. A separate Sample Submission Sheet should be filled out for this analysis

and SAC B021 is used. Reason Code 02 should be used on all reference station Sample Submission

Sheets.

Chesapeake Bay Nutrient Loading Stations

This station type is assigned either SAC 012 or 612 and all sites are within the Chesapeake Bay

watershed. SAC 612 stations are in geographic areas that could be underlain by Marcellus shale; this

SAC contains a few additional analytes typically associated with Marcellus formation water. Reason

Code 01 is used on the Sample Submission Sheet for the monthly, routine sample collected at these

Chesapeake Bay stations.

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Storm Sampling: Since these stations are also part of the CBP NTN, in addition to the monthly routine

sample, eight additional storm samples are collected throughout the sampling year. A storm event is

characterized with a rising discharge (cfs) of at least twice that of the pre-storm average daily

discharge. The assigned SAC, field parameters, and a suspended sediment sample is collected. These

samples are identified with Reason Code 014 on the datasheet. Ideally, one suspended sediment

concentration (SSC) and one SSC with sand and fine particle analyses should be collected per quarter.

If a storm event occurs on the scheduled routine sampling day, the sample is identified as “routine,

storm-impacted” and should be assigned Reason Code 015 on the datasheet. If feasible, an SSC

sample should be collected also. These “routine, storm-impacted” samples are identified as the routine

monthly sample and not as one of the eight storm samples. All suspended sediment samples are

shipped to the USGS Ohio-Kentucky-Indiana Water Science Center Sediment Laboratory in Louisville,

KY.

Storm sampling for Chesapeake Bay Nutrient Loading stations is the only targeted sampling in the

WQN. More detailed information on the sampling requirements for storm samples can be found in the

CBP’s Non-Tidal Water Quality Monitoring document (CBP 2008).

Lake Stations

Lake stations are assigned SAC 083. Lakes are additionally sampled for chlorophyll-a and pheophytin

under SAC B059. Lake Erie and Presque Isle Bay bimonthly samples are collected at mid-depth. For

all other lakes, two samples are collected from one site during mid-summer stratification conditions.

These sites correspond to the deepest point in each lake with one sample collected a meter below the

surface (top sample) and the second sample from one meter from the bottom (bottom sample). A water

quality profile, to include temperature, dissolved oxygen, pH, and specific conductance, is recorded at

this site through the vertical water column and an aliquot from the top sample is filtered for chlorophyll-

a analysis. Chlorophyll-a sampling is conducted according to Chlorophyll A Sampling Method in Lakes,

Ponds and Reservoirs (Lathrop 2017). Lake sampling protocols can be found in Evaluations of

Phosphorus Discharges to Lakes, Ponds, and Impoundments (DEP 2013). It is recommended that at

least one other site should be collected near the lake inlet. This site will follow the same procedures as

the deepest point site.

Pesticide Sampling

Due to the sampling design of the Water Quality Network, WQN stations are often utilized to gather

data for specific projects. Pesticide grab samples are collected between March 1st and September 30th

at eleven stations. Three of these are sampled monthly with two additional samples collected during

storm events. The remaining stations are sampled once during the March 1st through September 30th

sampling window. All samples are sent to the USGS National Water Quality Laboratory (NWQL) for

analysis. Samples will be delivered to the USGS for shipment within one day of sampling or will be

shipped directly to the NWQL within one day of sampling.

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POCIS Sampling

The WQN is also being utilized to monitor hormones and other endocrine-disrupting compounds

(EDCs). Polar Organic Chemical Integrative Samplers (POCIS) are a type of passive sampling device

being deployed at 21 WQN stations to collect water quality samples of low-level environmental

contaminants. POCIS sampling will follow the procedures outlined in the Passive Water Chemistry

Data Collection Protocol in Chapter 4 for the Monitoring Book (Shull and Lookenbill 2021). Analysis of

the wastewater indicators will be performed at the USGS National Water Quality Laboratory (NWQL) in

Denver, CO and the estrogenicity samples at the USGS Leetown Science Center in Kearneyville, WV.

BIOLOGICAL SAMPLING

All biological sampling is conducted using the protocols outlined in Chapter 3 of the Monitoring Book

(Shull and Lookenbill 2021). At a minimum, benthic macroinvertebrate data are collected at standard

stations every other year between August 1st and September 30th. Standard stations that are wadeable

in the non-summer months will be sampled between November 1st and April 30th. Macroinvertebrates

are collected annually at Chesapeake Bay stations between November 1st and April 30th. Reference

stations either rotate annually between a spring (March 1st to April 30th) and fall (Nov. 1st to Dec. 31st)

collection or are sampled twice per year (once in spring and once in fall).

Data collection per the wadeable riffle-run protocol is appropriate at most stations, however, WQN

stations can be sampled using any of the protocols outlined in Chapter 3 of the Monitoring Book (Shull

and Lookenbill 2021) if appropriate. The appropriate data collection protocols for each WQN is listed in

Subappendix D.3.

A physical habitat evaluation is conducted with every macroinvertebrate data collection effort. The

procedures in the Stream Habitat Data Collection Protocol within Chapter 5 of the Monitoring Book

(Shull and Lookenbill 2021) should be used. The majority of WQN stations will use the riffle/run habitat

parameters, however, a few stations will use the multihabitat parameters if they are determined to be

low gradient. Please contact the WQD if clarification is needed. Habitat field data sheets are available

in Appendix C in the Monitoring Book (Shull and Lookenbill 2021).

Qualitative plankton samples are collected annually at Lake Erie stations. Quantitative invertebrate or

plankton sampling and quantitative or qualitative fish sampling is optional at any station and may be

conducted at the discretion of the collector. Plankton data collection protocols are detailed in

Quantitative Plankton Sampling Method for Lakes (DEP 2015) and fish sampling protocols are in

Chapter 3 of the Monitoring Book (Shull and Lookenbill 2021).

Fish tissue is sampled periodically at the rate of approximately 100 stations per year following the

protocols outlined in Chapter 3 of the Monitoring Book (Shull and Lookenbill 2021). Tissue sampling

locations are determined annually during the first quarter of the calendar year preceding sample

analysis and rotated throughout the WQN to provide periodic but statewide coverage and to maintain

surveillance on problem waters. Fish tissue (fillets) is analyzed for appropriate priority pollutants to

D-12

assess its suitability for human consumption.

FLOW MEASUREMENT

Stream discharge (flow volume) is measured or calculated each time a water chemistry sample is

collected. USGS stream gaging facilities and/or extrapolation equations are utilized whenever possible.

Where no USGS facilities/equations exist, stream discharge is measured using U.S. Army Corps of

Engineers and private gaging facilities or calculated according to procedures outlined by USGS

(Turnipseed and Sauer 2010). Lake levels for Lake Erie and Presque Isle Bay stations are measured at

the U.S. Coast Guard station at the entrance to Erie Harbor.

Flow Measurement Procedures

The Pennsylvania WQN utilizes several types of stage gages to provide stream discharge. The two

dominant types are the Wire-Weight (WW) gage and the Electric Tape (ET) gage. Both are non-

recording gages that yield readings indirectly via measurement from a fixed point to the water surface.

Other gages that may be used include a Staff gage, Analog-to-Digital Recorder (ADR) dial, Strip Chart,

Hand Tape, Crest Stage Gage, Lock Chamber or Telemark (TMK). Descriptions on the various flow

measurement procedures are in Subappendix D.7.

The Water Resources Division of the USGS recommends that an outside gage such as a Staff gage or

a WW gage be used by DEP personnel. If no outside gage is present or it is unreadable, USGS prefers

a reading from the ADR dial or the ET gage, in that order. The base gages for the network stations are

primarily ET gages or WW gages. If neither of these types are present, a Staff gage may function as

the base gage for that site.

DATA AVAILABITY AND MANAGEMENT

All chemical data analyzed by DEP BOL are entered by BOL staff into DEP’s internal Oracle database.

The collector or DEP WQD staff are responsible for entering the field chemistry, station information,

comments, and USGS OH-KY-IN Sediment Lab results into the Oracle database. The data are

uploaded quarterly by DEP via the Water Quality Exchange (WQX) to the United States Environmental

Protection Agency’s (EPA) database. This data is made available to the public on the Water Quality

Portal (https://www.waterqualitydata.us). Macroinvertebrate samples are processed, and the taxa

identified and enumerated by a contractor or by WQD staff. The macroinvertebrate data are entered

into the WQD’s internal Streams and Lakes Integrated Management System (SLIMS) database.

Analyses of trends at selected WQN stations are published in DEP's Biennial Integrated Water Quality

Monitoring and Assessment Report.

QUALITY ASSURANCE

A quality assurance project plan has been developed to describe the procedures used to implement the

WQN in both the field and laboratory. Copies of this plan can be obtained from the Bureau of Clean

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Water’s WQD or EPA Region III. One sample "blank" is submitted bimonthly by each field office

involved in sample collection and duplicate or "split samples" are collected at the rate of one in every

20 routine WQN samples. Qualified WQD staff routinely audit WQN field staff to ensure sampling

consistency between staff and to evaluate the utility of each station. Additional quality assurance for

specific data collection protocols can be found throughout the Monitoring Book (Shull and Lookenbill

2021).

LITERATURE CITED

Barbour, M. T., J. Gerritsen, B. D. Snyder, and J. B. Stribling. 1999. Rapid bioassessment protocols for

use in streams and wadeable rivers: Periphyton, benthic macroinvertebrates and fish. 2nd

edition. EPA 841-B-99-002. U.S. Environmental Protection Agency, Office of Water,

Washington, D.C.

Buchanan, T. J., and W. P. Somers. 1969. Discharge measurements at gaging stations. Book 3,

chapter A8, page 65. in National field manual for the collection of water quality data: Techniques

and methods. U.S. Department of the Interior, U.S. Geological Survey, Reston, VA. (Available

online at: https://pubs.er.usgs.gov/publication/tm3A8.)

CPB. 2008. Non-tidal water quality monitoring. Chapter V in Non-Tidal Water Quality Field Procedures.

Chesapeake Bay Program, Annapolis, MD.

DEP. 2013. Evaluations of phosphorus discharges to lakes, ponds, and impoundments. Pennsylvania

Department of Environmental Protection, Harrisburg, PA.

DEP. 2015. Quantitative plankton sampling method for lakes (modified standard Method 1002a).

Pennsylvania Department of Environmental Protection, Harrisburg, PA.

Lathrop, B. F. 2017. Chlorophyll a sampling method in lakes, ponds and reservoirs. Pennsylvania

Department of Environmental Protection, Harrisburg, PA.

Lookenbill, M. J., and R. Whiteash (editors). 2021. Water quality monitoring protocols for streams and

rivers. PA Department of Environmental Protection, Harrisburg, PA.

Turnipseed, D. P. and V. B. Sauer. 2010. Discharge measurements at gaging stations: United States

Geologic Survey. Book 3, chapter A8. in National field manual for the collection of water quality

data: Techniques and methods. U.S. Department of the Interior, U.S. Geological Survey,

Reston, VA. (Available online at: https://pubs.usgs.gov/twri/twri3a8/.)

Wilde, F. D. (editor). 2004. Processing of water samples. Book 9, chapter A5 (version 2.2). in National

field manual for the collection of water quality data: Techniques and methods. U.S. Department

of the Interior, U.S. Geological Survey, Reston, VA. (Available online at:

https://www.usgs.gov/mission-areas/water-resources/science/national-field-manual-collection-

water-quality-data-nfm?qt-science center objects=0#qt-science center objects.)

USGS. 2006. Collection of water samples. Book 9, chapter A4 (version 2.0). in National field manual for

the collection of water quality data: Techniques and methods. U.S. Department of the Interior,

U.S. Geological Survey, Reston, VA. Available online at:

https://pubs.er.usgs.gov/publication/twri09A4.

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D.1 STANDARD ANALYSIS CODES (SAC)

D-15

WQN PARAMETER LISTS

1. ALL STATIONS (standard field analyses) pH Temperature Dissolved Oxygen Specific Conductance

** % Dissolved Oxygen and Turbidity should also be reported, if collected, using data collection protocols in Lookenbill and

Whiteash (2021).

2. STANDARD STATIONS (SAC 010)

Test Code Test Description Test Code Test Description 00410 Alkalinity (Alk) 01105H Aluminum, Total (Al)

00610A Ammonia Nitrogen (NH3 N) 99020 Bromide (Br)

00916A Calcium, Total (Ca) 00940 Chloride (Cl)

01042H Copper, Total (Cu) 00900 Hardness, Total (Hard)

01045A Iron, Total (Fe) 01051H Lead, Total (Pb)

00927A Magnesium, Total (Mg) 01055A Manganese, Total (Mn)

01067A Nickel, Total (Ni) 00620 Nitrate Nitrogen (NO3 N)

00615 Nitrite Nitrogen (NO2 N) 00600A Nitrogen, Total (N)

00403 pH, Lab (PH) 00665A Phosphorus, Total (P)

70507A Phosphorus Orthophosphate, Total P(O) 00937A Potassium, Total (K)

00929A Sodium, Total (Na) 00095 Specific Conductivity (SPC)

00945 Sulfate, Total (SO4) 70300U Total Dissolved Solids @ 180C (TDS)

00530 Total Suspended Solids (TSS) 01092A Zinc, Total (Zn)

3. CHESAPEAKE BAY NUTRIENT LOADING STATIONS (SAC 012)

Test Code Test Description Test Code Test Description

00410 Alkalinity (Alk) 01105A Aluminum, Total (Al)

00608A Ammonia Nitrogen, Dissolved (NH3 N) 00610A Ammonia Nitrogen, Total (NH3 N)

99020 Bromide (Br) 00916A Calcium, Total (Ca)

00940 Chloride (Cl) 01042H Copper, Total (Cu)

00900 Hardness, Total (Hard) 01045A Iron, Total (Fe)

01051H Lead, Total (Pb) 00927A Magnesium, Total (Mg)

01055A Manganese, Total (Mn) 01067A Nickel, Total (Ni)

00631A Nitrate & Nitrite, Dissolved (NO3 + NO2) 00630A Nitrate & Nitrite, Total (NO3 + NO2)

00602A Nitrogen, Dissolved (N) 00600A Nitrogen, Total (N)

00680 Organic Carbon, Total 00403 pH, Lab (PH)

00671A Phosphorus Orthophosphate, Dissolved P(O) 70507A Phosphorus Orthophosphate, Total P(O)

00666A Phosphorus, Dissolved (P) 00665A Phosphorus, Total (P)

00937A Potassium, Total (K) 00929A Sodium, Total (Na)

00095 Specific Conductivity (SPC) 00945 Sulfate, Total (SO4)

70300U Total Dissolved Solids @ 180C (TDS) 00530 Total Suspended Solids (TSS)

01092A Zinc, Total (Zn)

D-16

4. REFERENCE STATIONS (SAC 015 & B021*)

Test Code Test Description Test Code Test Description 70508

01106H

Acidity, Total

Aluminum, Dissolved (Al) 00410

01105H

Alkalinity (Alk)

Aluminum, Total (Al)

00608A Ammonia Nitrogen, Dissolved (NH3 N) 00610A Ammonia Nitrogen, Total (NH3 N)

01007H Barium, Total (Ba) 00314 Biochemical Oxygen Demand (CBOD)

01022K Boron, Total (B) 99020 Bromide (Br)

01025H Cadmium, Dissolved (Cd) 00915A Calcium, Dissolved (Ca)


01040H Copper, Dissolved (Cu) 01042H Copper, Total (Cu)

MMTECMF E. coli* 31616 Fecal Coliform* (FC)

00900 Hardness, Total (Hard) 01046A Iron, Dissolved (Fe)

01045A Iron, Total (Fe) 01130A Lithium, Dissolved (Li)

01132A Lithium, Total (Li) 01049H Lead, Dissolved (Pb)

01051H Lead, Total (Pb) 00925A Magnesium, Dissolved (Mg)

00927A Magnesium, Total (Mg) 01056H Manganese, Dissolved (Mn)

01055H Manganese, Total (Mn) 01065H Nickel, Dissolved (Ni)

01067H Nickel, Total (Ni) 00631A Nitrate + Nitrite N, Dissolved

00630A Nitrate + Nitrite N, Total 00600A Nitrogen, Total (N)

00681 Organic Carbon, Dissolved 00680 Organic Carbon, Total

82550 Osmotic Pressure (OP) 00403 pH, Lab (PH)

00666A Phosphorus, Dissolved (P) 00665A Phosphorus, Total (P)

00671A

00937A

Phosphorus Orthophosphate, Dissolved P

Potassium, Total (K) 70507A

01147H

Phosphorus Orthophosphate, Total P

Selenium, Total (Se)

00929A Sodium, Total (Na 00095 Specific Conductivity (SPC)

01082A Strontium, Total (Sr) 00945 Sulfate, Total (SO4)

70300U Total Dissolved Solids @ 180C (TDS) 00530 Total Suspended Solids (TSS)

01090H Zinc, Dissolved (Zn) 01092A Zinc, Total (Zn)

5. LOW ALKALINITY STATIONS (SAC 016)


70508 Acidity, Total 00410 Alkalinity (Alk)

01106H Aluminum, Dissolved (Al) 01105H Aluminum, Total (Al)

00610A Ammonia Nitrogen, Total (NH3 N) 00314 Biochemical Oxygen Demand (CBOD)

99020 Bromide (Br) 00915A Calcium, Dissolved (Ca)


01040H Copper, Dissolved (Cu) 01042H Copper, Total (Cu)

00900 Hardness, Total (Hard) 01046A Iron, Dissolved (Fe)

01045A Iron, Total (Fe) 01130A Lithium, Dissolved (Li)

01132A Lithium, Total (Li) 01049H Lead, Dissolved (Pb)

01051H Lead, Total (Pb) 00925A Magnesium, Dissolved (Mg)

00927A Magnesium, Total (Mg) 01056H Manganese, Dissolved (Mn)

01055H Manganese, Total (Mn) 01065H Nickel, Dissolved (Ni)

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01067H Nickel, Total (Ni) 00620 Nitrate Nitrogen (NO3 N)

00615 Nitrite Nitrogen (NO2 N) 00600A Nitrogen, Total (N)

00403 pH, Lab (PH) 00665A Phosphorus, Total (P)

70507A Phosphorus Orthophosphate, Total P(O) 00937A Potassium, Total (K)

00929A Sodium, Total (Na) 00095 Specific Conductivity (SPC)

00945 Sulfate, Total (SO4) 70300U Total Dissolved Solids @ 180C (TDS)

00530 Total Suspended Solids (TSS) 01090H Zinc, Dissolved (Zn)

01092H Zinc, Total (Zn)

6. ATMOSPHERIC DEPOSITION REFERENCE STATIONS (SAC 122): Includes B021 and all SAC 015 parameters plus:

Test Code 01106D

Test Description Aluminum, Dissolved 0.1 micron filter

Test Code 00420C

Test Description Carbon Dioxide, Total (CO2)

7. STANDARD TDS STATIONS (SAC 610): Includes all SAC 010 parameters plus:


01007H Barium, Total (Ba) 01022K Boron, Total (B)

01132A Lithium, Total (Li) 82550 Osmotic Pressure (OP)

01147H Selenium, Total (Se) 01082A Strontium, Total (Sr)

8. CHESAPEAKE BAY NUTRIENT LOADING TDS STATIONS (SAC 612): Includes all SAC 012 parameters plus:



01130A Lithium, Dissolved (Li) 01132A Lithium, Total (Li)

82550 Osmotic Pressure (OP) 01147H Selenium, Total (Se)

01082A Strontium, Total (Sr)

9. LOW ALKALINITY TDS STATIONS (SAC 616): Includes all SAC 016 parameters plus:



82550 Osmotic Pressure (OP) 01147H Selenium, Total (Se)

01082A Strontium, Total (Sr)

10. RADIOLOGICAL (Suite RAD99): Alpha and Beta may be listed on the Sample Submission Sheet as

additional parameters, for any WQN station, to supplement the selected analysis code. DEP Water Quality

Division staff must be aware of these additions.

11. LAKE STATIONS (SAC 083 & B059*)


00410 Alkalinity, Total (Alk) 01105H Aluminum, Total (Al)

00610A Ammonia Nitrogen, Total (NH3 N) 01002H Arsenic, Total (As)

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01007A

71870

Barium, Total (Ba)

Bromide (Br)

01022K

00916A

Boron, Total (B)

Calcium, Total (Ca) 00940

01042H

Chloride (Cl)

Copper, Total (Cu)

962*

00900

Chlorophyll a (fluorometric)

Hardness, Total (Hard)

01045A Iron, Total (Fe) 01051H Lead, Total (Pb)

00927A Magnesium, Total (Mg) 01055H Manganese, Total (Mn)

71900I

00600A

PHEOPHY*

01147H

Mercury, Total

Nitrogen, Total (N)

Pheophytin

Selenium, Total (Se)

00620A

00403

00665A

00929A

Nitrate Nitrogen (NO3 N)

pH, Lab (PH)

Phosphorus, Total (P)

Sodium, Total (Na)

00095 Specific Conductivity (SPC) 01082A Strontium, Total (Sr)

00945

00530

01092H

Sulfate, Total (SO4)

Total Suspended Solids (TSS)

Zinc, Total (Zn)

70300U

82079

Total Dissolved Solids @ 180C

Turbidity, Nephelmetric

12. FISH TISSUE STATIONS

a. SAC 059


71946 Barium, Total (Ba) 71940 Cadmium, Total (Cd)

71939 Chromium, Total (Cr) 71937 Copper, Total (Cu)

71936 Lead, Total (Pb) 71930 Mercury, Total (Hg)

71945 Selenium, Total (Se) 71947 Strontium, Total (Sr)

b. PCBs (SAC PCBF)

CAS # Test Description CAS # Test Description

11104282 Arochlor 1221 11141165 Arochlor 1232



c. PESTICIDES (SAC PESTF)

CAS # Test Description CAS # Test Description

72548 4,4-DDD 72559 4,4-DDE

50293 4,4-DDT 309002 Aldrin

319846 Alpha-BHC 5103719 Alpha-Chlordane

3734483 Chlordene 5103731 Cis-Nonachlor

60571 Dieldrin 72208 Endrin

5103742 Gamma-Chlordane 58899 Gamma-BHC (Lindane)

76448 Heptachlor 1024573 Heptachlor Epoxide

72435 Methoxychlor 2385855 Mirex

53190 O,P-DDD 3424826 O,P-DDE

789026 O,P-DDT 26880488 Oxychlordane

39765805 Trans-Nonachlor

D-19

D.2 ACTIVE WATER CHEMISTRY STATIONS

D-31

D.3 ACTIVE BENTHIC MACROINVERTEBRATE STATIONS

D-44

D.4 DISCONTINUED STATIONS

D-58

D.5 WQN OBJECTIVES

D-59



WATER QUALITY NETWORK OBJECTIVES

2020

D-60

Prepared by:

Molly Pulket and Heidi Biggs

Pennsylvania Department of Environmental Protection





2019

Edited by:

Molly Pulket






2020

D-61

OBJECTIVES OF THE WATER QUALITY NETWORK

The Pennsylvania Water Quality Network (WQN) is a statewide, fixed-station sampling network that

has been gathering important surface water quality data across the Commonwealth for decades. The

design of the network, the design of sampling plans in the network, and the decades of data available

at some stations have allowed WQN data to be used for many purposes by the Pennsylvania

Department of Environmental Protection (DEP), other state and federal agencies, and other interested

parties. This document describes the key uses of WQN water chemistry and macroinvertebrate data.

For more information on this sampling network, please visit the WQN webpage at DEP Water Quality

Network.

Trend Analysis

Trend analysis is one of the core functions of the WQN. The WQN’s fixed-station monitoring strategy is

uniquely suited for measuring changes in water quality through time. Trend data collected in the WQN

is used to monitor incremental progress in water quality, to calibrate Total Maximum Daily Load (TMDL)

models, and to track sources and timing of potential impacts to water quality. Analyses of trend data

are also one of the most efficient and accessible ways to communicate vast amounts of water quality

data to a variety of stakeholders, including the public, which promotes transparency.

Example

As part of a joint effort, DEP partners with the Susquehanna River Basin Commission (SRBC), the

United States Geological Survey (USGS), and the Chesapeake Bay Program (CBP) in a rigorous

sampling program to measure nutrient and sediment concentrations at strategic locations within the

Susquehanna River basin to better understand nutrient and sediment inputs to the Chesapeake Bay.

This monitoring network is incorporated into the WQN and consists of six mainstem river stations and

thirty tributary stations. From this extended WQN program, SRBC reports on trends in nutrient and

suspended sediment concentrations, loads, and yields in the Susquehanna River basin. Information

from these reports helps managers from multiple agencies create and adapt appropriate action plans.

Permitting

Data from the WQN informs the evaluation and development of water quality-based effluent limitations

(WQBEL) in National Pollutant Discharge Elimination System (NPDES) permits which control the

discharge of pollutants from point sources into surface waters. WQBELs, as defined in 25 Pa. Code

section 96.1, are effluent limitations based on the need to maintain or attain the water quality criteria

and to assure protection of existing and designated uses. DEP often uses WQN data as background

data for naturally occurring pollutants when determining whether a stream has assimilative capacity to

accept a pollutant limit and in developing WQBELs where there is reasonable potential to exceed water

quality criteria.

WQN data also provide important background data when permitting discharges in High Quality Waters

(HQ) and Exceptional Value Waters (EV) where existing water quality is protected and maintained. In

permitting discharges to these special protection waters, data from a WQN station near the proposed

D-62

or existing discharge site, or from a WQN station that is representative of the proposed or existing

discharge site, can be used to develop non-degrading effluent limitations.

Additionally, since the toxicity of several pollutants vary with hardness (e.g., copper, lead, nickel, silver,

and other metals) or pH (e.g., ammonia), many Chapter 93 water quality criteria require background

hardness or pH to use in calculating criteria values. In these situations, WQN data can be used to

eliminate the use of default hardness or pH values, or the need for additional data collection to

calculate appropriate water quality criteria values.

Example

DEP permit writers can use WQN data to develop non-degrading effluent permit limits for discharges to

HQ and EV streams. A DEP permit writer used water chemistry data collected at WQN station 195 on

Rock Run to calculate the non-degrading effluent limitations for an Exceptional Value tributary to

French Creek in Chester County. The WQN station allowed the permit writer to use data that is

representative of the watershed containing the permitted discharge, instead of default data values or

requiring the need to collect additional data.

Water Quality Assessments for Federal Clean Water Act Sections 303(d) and 305(b)

The WQN is crucial for Pennsylvania to meet federal mandates to monitor water quality and to assess

monitored water quality against water quality standards. Depending on the type of WQN station, certain

chemical parameters are collected frequently enough to determine if water quality criteria are being

met. This information is one part of the overall surface water monitoring program that helps build the

Pennsylvania Integrated Water Quality Monitoring and Assessment Report, which is required by the

federal Clean Water Act on a biennial basis. In addition, biological assessment determinations can also

be made at WQN stations where biological data is collected using current assessment methodologies.

Example

Historical and current data for nitrates collected at the WQN station on Connoquenessing Creek

located downstream of an industrial discharge in Butler County was instrumental in documenting the

impairment and subsequent restoration of the creek’s Potable Water Supply (PWS) use. The PWS use

of Connoquenessing Creek downstream of the discharge was listed as impaired on the 303(d) list for

exceeding the nitrite plus nitrate PWS criterion in 25 Pa. Code § 93.7 as documented in historical WQN

data. After the industrial discharger discontinued nitric acid pickling, WQN data documented water

quality in Connoquenessing Creek downstream of the discharge as no longer exceeding the nitrite plus

nitrate PWS criterion. In this instance, WQN data was central to both the listing and subsequent

delisting of Connoquenessing Creek.

Data to Inform Permits

The WQN is specifically designed to evaluate both the quality of Pennsylvania's surface waters and the

effectiveness of DEP’s water quality management program. Identifying changes in water quality at

certain locations over time helps ensure permits are written accurately and are protective of water

quality standards. Additionally, the WQN serves not only to help inform permitting, but also help

operators describe in permit applications how they are able to comply with water quality standards.

D-63

Example

The WQN data was essential in helping show compliance with water quality standards as discussed in

the industrial discharge example above. Although the WQN data initially documented the impairment,

WQN data also helped the operator demonstrate compliance with water quality standards after the

process change at the plant.

Threatened and Endangered Species Protection

Threatened and endangered species are an ongoing concern related to NPDES discharges. The

United States Fish and Wildlife Service has recommended stringent criteria for multiple pollutants

including ammonia-nitrogen, chloride, TDS, and nickel to protect threatened and endangered mussel

species. As noted in the permitting discussion above, WQN data is essential in accurately evaluating

appropriate criteria for some of these pollutants since the availability of background data can have a

significant impact on these evaluations. Not only is it critical to have accurate background data of the

pollutants of concern but it is also critical to have accurate hardness and pH background data to

calculate appropriate pollutant criteria and associated potential impact areas.

Example

Endangered mussels are of special concern in the Allegheny River, French Creek, Shenango River,

and many tributaries to these surface waters. Segments of all three main streams have been

designated as “critical habitat” for the rabbitsfoot mussel, and many other threatened and endangered

mussel species. Numerous NPDES permits in this region of Pennsylvania have been issued with

concurrence by US Fish and Wildlife Service as protective of endangered species, using a permitting

strategy that includes a robust evaluation of WQN data.

Emerging Science

Due to the years- to decades-long historical records of physical, chemical, and biological data collected

at many stations, WQN sites often serve as prime locations to collect new data (e.g., contaminants of

emerging concern) and to develop new collection protocols (e.g., periphyton monitoring protocols).

The ability to contextualize new kinds of data in robust historical records characterizing particular sites

across a range of parameters and assemblages can greatly reduce the cost and time needed to

expand monitoring approaches when novel environmental concerns arise.

Example

During an intensive study of the Susquehanna River, WQN stations were almost exclusively used when

selecting locations to place passive chemical samplers designed to detect hundreds of contaminants of

emerging concern. The combination of this new data and the existing WQN data substantially reduced

the overall cost of the project, compared with project costs if the passive samplers were deployed in

locations without robust water quality network data.

D-64

D.6 FIELD PREPARATION AND FIXATIVE CHART

D-65

Standard Analysis Code

(SAC) Parameters Bottle Ware Field Prep/Fixative

010

general chemistry 1 - 500 ml HDPE plastic bottle iced to 4°C

N/P 1 - 500 ml HDPE plastic bottle 10% H2SO4 to pH <2; iced to 4°C

metals, total 1 - 125 ml HDPE plastic bottle 1:1 HNO3 to pH <2; iced to 4°C

012


general chemistry, dissolved 1 - 125 ml HDPE plastic bottle field filtered; iced to 4°C


N/P, dissolved 1 - 125 ml HDPE plastic bottle field filtered; 10% H2SO4 to pH <2; iced to 4°C


TOC 2 - 40 ml amber VOA vials 10% H2SO4 to pH <2; iced to 4°C

015

general chemistry plus CBOD 3 - 500 ml HDPE plastic bottle iced to 4°C





metals, dissolved 1 - 125 ml HDPE plastic bottle field filtered; 1:1 HNO3 to pH <2; iced to 4°C

Organic Carbon, Total 2 - 40 ml amber VOA vials 10% H2SO4 to pH <2; iced to 4°C

Organic Carbon, Dissolved 2 - 40 ml amber VOA vials field filtered: 10% H2SO4 to pH <2; iced to 4°C

016





083




122






D-66


Organic Carbon, Total 2 - 40 ml amber VOA vials 10% H2SO4 to pH <2; iced to 4°C

Organic Carbon, Dissolved 2 - 40 ml amber VOA vials field filtered: 10% H2SO4 to pH <2; iced to 4°C

aluminum, monomeric (R01106D) 1 - 125 ml HDPE plastic bottle field filtered (0.1 um); 1:1 HNO3 to pH <2; iced to 4°C

610 *all SAC010 bottles plus






B021 fecal coliform, E.coli 1 - 125 ml plastic bottle sterilized & pretreated with sodium thiosulfate

iced to ≤ 6°C

RAD99 gross alpha, gross beta 2 - 500 ml glass bottle iced to 4°C

B029 chlorophyll a 1 - 47 mm glass fiber filter in

petri dish wrap in aluminum foil and place on dry ice or ice to 4°C &

freeze within 24 hrs

-- suspended sediment 1 - 500 ml plastic bottle store in cool dark place

All WQN stations require the following STANDARD FIELD ANALYSES: Temp (°C), pH, Dissolved Oxygen, and Specific Conductance. These results should be recorded on the Sample Submission Sheet.

* Must be shipped to USGS lab in Louisville, KY.

D-67

D.7 STREAM FLOW MEASUREMENT PROTOCOLS

D-68

Wire-Weight Gages

A wire-weight (WW) gage consists of a drum wound with a single layer of cable, a bronze weight

attached to the end of the cable, a graduated disc and counter, and a check bar of known elevation.

The gage is housed within a small cast aluminum lockable box and generally mounted to a rigid

structure (i.e., bridge or trestle) directly above the water surface. The graduated disc is permanently

connected to the counter and to the shaft of the drum. The drum reel is equipped with pawl and ratchet

for holding the suspended weight at any desired elevation.

The check-bar for the WW gage is set by levels as a reference elevation to gate datum. The gage is set

so that when the bottom of the suspended weight is at the water surface, the gage height is indicated

by the combined readings of the counter and the graduated disc.

Reading Instructions

• Unlock the case and gently open the cover. If the cover jams and will not open freely, do not

force it open. The cover has probably become jammed against the crank handle. Try to move

the handle into a position that will allow the cover to open. At this point, make sure the pawl is

engaged or that you can hold onto the handle to prevent the weight falling to the stream bed

and unspooling the cable.

• Inspect the cable. It should be tightly wound with windings that touch each other, with no

overlaps. If cable is not properly wound, check-bar and water-surface readings will be

incorrect. If necessary, unwind the cable and utilizing the level-wind pulley and gloves, rewind

the cable onto the reel. (This should only be necessary in extreme situations.)

• The check-bar is found directly under the weight as it hangs in the box and has two positions,

the forward position (away from you) which prevents the weight from falling to the bottom of

the box while locked (and into the stream when opened), and a rear position (toward you)

which allows the weight to be lowered to the stream. You should find the check-bar in the

forward position. Elevations are run to the check-bar in the forward position and the gage is set

to read this elevation when the weight just makes contact with the check-bar.

• In order to determine the check-bar reading,

o Make sure the check-bar is in the forward position.

o Lower the weight until the weight just lightly touches the check-bar.

o Record this reading to three decimal places. This will involve estimating the last digit.

• Raise the weight and slide the check-bar to the rear position. Lower the weight until it just

lightly touches the surface of the water and record your reading to two decimal places.

Average the peaks and troughs of elevation if the water surface is surging. You should repeat

this process at least once. If the air temperature is below freezing, the first time the weight

touches the water, a layer of ice will form on the bottom of the weight causing it to become

slightly longer. Repeated contacts with the water surface continues to add layers of ice to the

bottom of the weight. For this reason, water surface elevations should be made as accurately

as possible on the first or second try.

• Engage the pawl (this will prevent the weight from falling and unspooling the cable should the

crank slip from your hand while rewinding) and crank the weight to its original position. Slide

D-69

the check-bar to its forward notch. The crank handle should be located in the rear position,

allowing the cover to close without touching the crank handle.

• Close the cover and lock the case.

Electric Tape Gages

The electric tape (ET) gage consists of a steel tape graduated in feet and hundredths, to which is

fastened a cylindrical weight (usually brass), a reel in a frame for the tape, a 4½-volt battery and a

voltmeter. One terminal of the battery is attached to a ground connection and the other to one terminal

of the voltmeter. The other terminal of the voltmeter is connected through the frame, reel and tape, to

the weight. The weight is lowered until it contacts the water surface. This contact completes the electric

circuit and produces a signal on the voltmeter. With the weight held in the position of first contact, the

water surface elevation reading is observed at the tape index (a black line indented just below the

voltmeter and usually immediately above the shelf) provided on the reel mounting. ET gages are

installed in a manner that will produce a dependable circuit, yielding a strong signal when the weight

touches the water surface.

Reading Instructions

The dog on the left side of the reel is disengaged by lifting the rear portion in an upward direction while

holding onto the handle on the right side. This allows the weight and tape to be lowered until it contacts

the water surface. This contact completes the electric circuit and produces a fluctuation on the

voltmeter on the front of the tape reel frame. Once a contact with the water surface has been made, the

weight is raised slightly until contact is broken, and then lowered very slowly until the water surface is

again contacted. This procedure is repeated until a consistent reading at the index line is produced.

This reading may be confirmed visually by shining a flashlight into the well at the spot where the weight

will make contact with the surface of the water and lowering the weight very slowly until contact with the

surface is visually confirmed (the voltmeter needle should also deflect at this time if working correctly).

The reading of the water surface elevation is then taken by observing the reading of the steel tape at

the index line on the gage. NOTE: HAZARD!! In order to make a visual observation of the weight

touching the water surface in the well, it is often necessary to raise the door (located on the floor) to the

well. Once the door is opened, extreme care should be exercised to ensure that nothing is dropped into

the well. This type of reading is best left to experienced personnel.

Possible Interferences

• If the weight touches any grounded item (steel piping or the steel side of an oil tube or a steel

pipe well) the circuit will be completed, and the voltmeter will deflect as if the water surface

has been contacted.

Solution: Physically move the tape past any items in the well that will produce a false reading.

Keep in mind that the tape should be kept as vertical as possible for accurate readings.

• If any oil has seeped into the well onto the water surface from the oil tube, the needle will not

deflect upon hitting the oil, but will deflect when weight is lowered through the oil and contacts

the water surface (this will not be an accurate water elevation).

Solution:

D-70

o Visually determine elevation of the top of the oil surface using the procedure described

above for visual readings and record this number.

o From the top of the oil very slowly lower the weight until a deflection of the voltmeter

occurs. This is the top of the water surface.

o Subtract this reading from the one made in step 1. and multiply the answer by 0.2.

o Subtract this value from the top of oil reading determined in step 1. This is the actual

surface elevation.

Example: 5.28 surface of oil

5.23 surface of water

0.05 difference

X.20 = .0100

5.28-.01 = 5.27 (actual water surface elevation)

• If the bottom of the weight is corroded, even slightly, the meter will not deflect until the water

in the well encounters a portion of the weight free of corrosion, completing the circuit. This

produces an erroneous reading as the circuit is not completed at the bottom of the weight, but

at some point along the side.

Solution: Using a piece of emery paper rub lightly over the bottom of the weight to remove

corrosion. It is also acceptable to lightly scrape the edge of a knife across the bottom of the

weight. Care must be taken to avoid removing any brass from the bottom of the weight as

repeated cleanings over years may change the weight length.

• If the bottom of the weight is corroded, even slightly, the meter will not deflect until the water

in the well encounters a portion of the weight free of corrosion, completing the circuit. This

produces an erroneous reading as the circuit is not completed at the bottom of the weight, but

at some point along the side.

Solution: Using a piece of emery paper rub lightly over the bottom of the weight to remove

corrosion. It is also acceptable to lightly scrape the edge of a knife across the bottom of the

weight. Care must be taken to avoid removing any brass from the bottom of the weight as

repeated cleanings over years may change the weight length.

• The cleaning of the weight is a delicate procedure as the weight must first be lowered a few

feet, so it may be grasped through the shelf door or opening. Very often the tape may be very

rusted at the point the tape enters the weight or just below that point. "Any twisting or bending

will break the tape off the weight." Should this happen, lay the weight on the shelf and

contact the USGS as soon as possible.

• Ice in the well.

Solution: Ice must be broken loose and, in some cases, removed. This is a solution that is

best left to experienced USGS technicians, and in no instance should DEP personnel enter

the well.

Staff Gages

Staff gages are simply rods or boards that are securely attached to a backing or permanent foundation.

These non-recording gages are graduated, and stage can be directly read. Vertical staff gages are

D-71

used as reference gages within stilling wells, and both vertical and inclined staff gages may be used as

outside reference gages. Lake height measurements for the WQN are obtained from staff gages.

Digital Recorders

Digital recorders are slow speed, battery operated instruments which record information in the binary

decimal code form by use of a paper-tape punch mechanism. Sixteen channel paper-tape is punched

with a four-digit number at preselected time intervals of 5, 15, 30 or 60 minutes. Stage heights of zero

to 99.99 feet can be recorded in increments of hundredths of a foot. A battery powers the sequencing

camshaft and electro-mechanical timing devices. Pins on the punch block are forced against the paper-

tape and code discs by spring action thereby recording the position of the discs at that moment. After

the paper-tape is punched, it is advanced, and the punch spring is compressed for the next readout

cycle. The tape can be either manually read to determine stage, or electronic translators can then

convert the punch-tape records to a magnetic tape form that is utilized to compute daily mean gage

height and daily stream discharge via digital computers.

Manual Measurement

A full stream width transect is required to manually measure stream flows. Total stream flow must be

calculated across this transect using the procedure of Buchanan and Somers (1969) described briefly

below. Record the depths and velocities for each of a minimum of 20 equally spaced points along the

transect, thereby constructing a stream channel profile and accompanying current velocity

measurements. Sum the products of each cell area calculation multiplied by velocity to determine total

flow (cfs) across the transect.

Telemark

Some gages are equipped with telecommunication capabilities that provide for their remote reading via

the internet.

E-1

APPENDIX E: QUALITY ASSURANCE PROJECT PLAN (QAPP): DATA COLLECTION FOR

THE PURPOSE OF EXISTING USE EVLUATIONS, PROTECTED USE ASSESSMENT, AND

EVALUATIONS OF POINT AND NONPOINT SOURCE IMPACTS TO SURFACE WATER

E-2

Prepared by:

Josh Lookenbill and Mark Hoger






2021

Disclaimer:

Appendix E is a copy of a QAPP approved by EPA in July 2021. An official, signed copy of the QAPP is

available upon request.

Project 2021: Document QA # 210096.2 Revision 02 July 2021

E-3

PENNSYLVANIA DEPARTMENT OF ENVIRONMENTAL PROTECTION

Bureau of Clean Water (BCW)

Water Quality Division (WQD)

SECTION A: PROJECT MANAGEMENT

A1. Title Page and Approval Sheet

TITLE: Quality Assurance Project Plan (QAPP): Data collection for

the purpose of existing use evaluations, protected use

assessments, and evaluations of point and nonpoint source

impacts to surface waters

EFFECTIVE DATE: August 1, 2021

AUTHORITY: Pennsylvania Clean Streams Law, 35 P.S. §§ 691.1 et. seq.,

the Federal Clean Water Act, 33 U.S.C.A §§ 1251 et. seq.,

25 Pa. Code Chapters 93 and 96.

POLICY: This guidance provides the established QAPP for water

quality data collection in surface waters of Pennsylvania

PURPOSE: The guidance was developed to establish and standardize

the Department of Environmental Protection’s (DEP) QAPP

procedures for data collection for existing use evaluations,

protected use assessments, and evaluations of point and

nonpoint source impacts to surface waters.

DISCLAIMER: The policies and procedures outlined in this guidance

document are intended to supplement existing requirements.

Nothing in the policies or procedures shall affect regulatory

requirements.

The policies and procedures herein are not an adjudication

or a regulation. There is no intent on the part of DEP to give

these rules that weight or deference. This document

establishes the framework within which DEP will exercise its

administrative discretion in the future. DEP reserves the


E-4

discretion to deviate from this policy statement if

circumstances warrant.

PAGE LENGTH: 37 Pages

DEP Project Manager’s Signature:

DEP Project Manager: Michael (Josh) Lookenbill

DEP Project Quality Assurance Manager’s Signature:

DEP Project Quality Assurance Manager: Mark Hoger

EPA Project Manager’s Signature: Note: This approval action represents EPA’s determination that the document(s) under review comply with

applicable requirements of the EPA Region 3 Quality Management Plan

[https://www.epa.gov/sites/production/files/2020-06/documents/r3qmp-final-r3-signatures-

2020.pdf] and other applicable requirements in EPA quality regulations and policies

[https://www.epa.gov/quality]. This approval action does not represent EPA’s verification of

the accuracy or completeness of document(s) under review, and is not intended to constitute

EPA direction of work by contractors, grantees or subgrantees, or other non-EPA parties.

EPA Project Manager: Steve Hohman

EPA Project Quality Assurance Manager’s Signature: Note: This approval action represents EPA’s determination that the document(s) under review comply with

applicable requirements of the EPA Region 3 Quality Management Plan

[https://www.epa.gov/sites/production/files/2020-06/documents/r3qmp-final-r3-signatures-

2020.pdf] and other applicable requirements in EPA quality regulations and policies

[https://www.epa.gov/quality]. This approval action does not represent EPA’s verification of

the accuracy or completeness of document(s) under review, and is not intended to constitute

EPA direction of work by contractors, grantees or subgrantees, or other non-EPA parties.

EPA Project Quality Assurance Officer: Elizabeth Gaige

Last Revised July 2021


E-5

A2. Table of Contents

Section A: Project Management ................................................................................... E-3

A1. Title Page and Approval sheet ........................................................................... E-3

A2. Table of Contents ............................................................................................... E-5

A3. Distribution List .................................................................................................. E-7

A4. Project Organization and Task Responsibility ................................................... E-8

A5. Problem Background ....................................................................................... E-10

A6. Project Task Description .................................................................................. E-12

A7. Quality Objectives and Criteria ........................................................................ E-14

A8. Training and Certifications ............................................................................... E-16

A9. Documents and Records ................................................................................. E-17

Section B: Data Generation and Acquisition .............................................................. E-22

B1. Experimental Design ........................................................................................ E-22

B2. Sampling Protocols .......................................................................................... E-24

B3. Sample Handling and Custody ........................................................................ E-26

B4. Analytical methods .......................................................................................... E-28

B5. Quality Control ................................................................................................. E-28

B6. Equipment Maintenance and Inspection .......................................................... E-30

B7. Instrument Calibration ...................................................................................... E-31

B8. Inspection/Acceptance of Supplies and Consumables .................................... E-31

B9. Non-Direct Measurements ............................................................................... E-32

B10. Data Management ......................................................................................... E-32

Section C: Assessment and Oversight ....................................................................... E-33

C1. Assessments and Response Actions .............................................................. E-33

C2. Reports to Management .................................................................................. E-33

Section D: Data Validation and Usability .................................................................... E-34

D1. Data Review, Verification and Validation ......................................................... E-34

D2. Verification and Validation of Methods ............................................................ E-34

D3. Reconciliation with User Requirements ........................................................... E-35

Literature Cited ........................................................................................................... E-36


E-6

E.1: Benthic Macroinvertebrate Subsampling and Taxonomic Identification Quality

Control ........................................................................................................................ E-39

Introduction ............................................................................................................. E-40

QC – Benthic Macroinvertebrate Subsampling ....................................................... E-40

QC – Benthic Macroinvertebrate Identification ........................................................ E-42

QC – Fish Identification ........................................................................................... E-43

Literature Cited ....................................................................................................... E-44


E-7

A3. Distribution List

This QAPP will be appended to the latest edition of the DEP Water Quality Monitoring

Protocols for Stream and Rivers (Lookenbill and Whiteash 2021), which is available on

the DEP Water Quality Division website at:

https://www.dep.pa.gov/Business/Water/CleanWater/WaterQuality/Pages/Data-

Collection-Protocols.aspx.

It will also be placed electronically in in-house records at the DEP. It can be made

available upon request. QAPP revisions will also be re-appended to DEP monitoring

protocols. In addition, this QAPP will be distributed to each of the DEP Regions and

cooperating programs who are involved with surface water and data collection. This will

ensure data will be collected according to the quality addressed in this QAPP. Any

revisions to this document will also be sent to those on the distribution list. The following

is the distribution list of current staff that oversees coordination in each region and each

respective agency:

• DEP Northwest Region – Joseph Brancato, Aquatic Biologist Supervisor

• DEP Southwest Region – Richard Spear, Aquatic Biologist Supervisor

• DEP Northcentral Region – Anne Hughes, Environmental Group Manager

• DEP Southcentral Region – Kristen Bardell, Aquatic Biologist Supervisor

• DEP Northeast Region – Walter Holtsmaster, Aquatic Biologist Supervisor

• DEP Southeast Region – Steven Unger, Aquatic Biologist Supervisor

• DEP Central Office – Josh Lookenbill, Monitoring Section Chief

• Department of Natural Resources and Conservation (DCNR) – Nick Decker,

Resource Manager

• Bureau of Laboratories (BOL) – Pamela Higgins, Special Assistant to Laboratory

Operations


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A4. Project Organization and Task Responsibility

The following is a list of key project personnel and their corresponding responsibilities.

Due to staff turnover, the list provided gives a general description of those involved. See

Figure 1 – Organizational Chart to better define their relationships.

• Sampling Operations – WQD Central Office Biologists, DEP Regional Biologists

• Sampling Quality Control (QC) – DEP Project Manager

• Laboratory Analysis – DEP Bureau of Laboratories (BOL), Outside laboratories

as needed

o Inorganic Division – Trace Metals & Sample Receiving Section and

Automated Analysis & Biochemistry Analytical Section

o Organic Division – Organic Chemistry Section

o Biological Services Division – Microbiology Lab, Giardia Lab and Microbial

Source Tracking Lab

• Laboratory Quality Control – BOL Quality Assurance and Safety Section

• Data and QC Processing – DEP Project Manager, WQD Central Office

Biologists, DEP Regional Biologists

• Data Review and Review QC – Falls to staff responsible for survey, could include

WQD Central Office Biologist or DEP Regional Biologist

• Performance and Systems Auditing – WQD, Monitoring Section and Assessment

Section

• Overall Project Coordination and QA – WQD, Monitoring Section

• Maintenance and Distribution of Official Approved QAPP – DEP Project Quality

Assurance Manager


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Figure 1. Organizational Chart


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A5. Problem Background

The primary purpose for monitoring surface waters is to assess and evaluate protected

uses and to broadly evaluate point and nonpoint sources and associated impacts on

surface waters in accordance with 25 Pa. Code §§ 93 and 96 of DEP’s regulations.

Monitoring is driven by the following primary objectives:

Assessment of Protected Water Uses

To investigate and determine attainment of water quality standards (WQS) and possible

sources and causes of impairment from point or non-point sources of conventional

pollutants and known or suspected water quality problems in streams, rivers and lakes.

These surveys are performed to reevaluate existing assessments and identify additional

sources and causes of water quality impairments identified by previous assessment

efforts. Chemical and bacteriological data is typically compared to water quality criteria

while considering frequency and duration with which a given criterion was developed

according to supporting technical documentation and assessment methods.

• Pennsylvania surface water quality criteria can be found in 25 Pa. Code § 93

(https://www.pacodeandbulletin.gov/Display/pacode?file=/secure/pacode/data/02

5/chapter93/chap93toc.html)

• Technical documentation

(https://www.dep.pa.gov/Business/Water/CleanWater/WaterQuality/Pages/Macro

invertebrates.aspx)

• Assessment methods

(https://www.dep.pa.gov/Business/Water/CleanWater/WaterQuality/Pages/Asses

sment-Methodology.aspx)

Biological data (macroinvertebrates and fish) is used to calculate indices that are

compared to attainment thresholds described in the assessment methods (Lookenbill

and Whiteash 2021) and accompanying technical documentation.

Data Usage

Data are used for generating the integrated report, listing impaired waterbodies as

required by Section 303(d), and to support the compliance and permitting programs by

defining the impact of specific discharges or land-based activities on receiving waters.

Physical, chemical and/or biological data collected during surveys are generally

evaluated using non-parametric, classification type analyses designed to display

differences or similarities between sampling stations and metric thresholds.


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Stream Redesignation and Existing Use Evaluations

To review and revise, if necessary, WQS to ensure that protected uses are accurate.

Data Usage

Data will be used to evaluate the need for antidegradation protection in selected

Pennsylvania waters. The evaluation will consider biotic and abiotic components of the

watershed ecosystem. Features such as water quality sensitivity, land use activities,

and the environmental and ecological significance of the watershed are evaluated in an

effort to determine the correct protected use per the requirements in 25 Pa. Code §

93.4b.

Maintenance of fixed station Water Quality Network (WQN)

To establish and maintain a database to assist in a) describing water quality conditions

across Pennsylvania streams, rivers and lakes; b) identifying long-term trends of

improvement or degradation in major waterbodies; c) determining the suitability of

waterbodies for water supply, recreation, fish, aquatic life, and other uses; d) providing a

limited measure of “background” water quality throughout Pennsylvania’s Ecoregions;

and e) providing a basis for evaluating success of the water quality management

program.

Data Usage

WQN data will be used to generate basic statistics to generally characterize water

quality by station; these values can then be compared to established standards and

criteria to determine the effectiveness of water quality management programs.

Furthermore, seasonally adjusted Kendall tests are applied to historic WQN data to

assess long-term trends in water quality using a Weighted Regressions on Time,

Discharge, and Season (WRTDS) model (Hirsch et al. 2010). Additionally, collection of

data at “ambient” stations provides a measure of background conditions at unaffected or

minimally affected sampling locations. These data provide both a measure of long-term

water quality trends resulting from airborne contaminants and background water quality

as identified by ecoregion. This information can then be used to develop permits by the

permitting programs or to determine the potential use attainment of nearby streams

degraded by anthropogenic inputs.

Other Objectives

To include cause and effect monitoring, point of first use determinations and protocol,

method and standards development.

Date Usage

Cause and effect monitoring data will be employed to investigate possible relationships


E-12

between point or nonpoint sources of conventional pollutants and known or suspected

water quality problems through the collection and analysis of biological, physical, and

chemical data. These surveys are designed primarily to monitor the effectiveness of

permitted treatment facilities but are also used to investigate reported or suspected

water quality impacts for nonpoint source and other pollution sources.

Point of first use surveys are completed as part of the evaluation and permitting of

wastewater discharges to intermittent and ephemeral streams, drainage channels and

swales, and storm sewer. The point of first surface water use establishes where

Chapter 93 WQS must be attained.

Protocol, method and standard development data collections will be used to implement

the Pennsylvania Clean Streams Law and the Federal Clean Water Act. To achieve

this, at a statewide level, WQS are developed to establish the protected uses, criteria,

and antidegradation requirements that surface waters must meet.

A6. Project Task Description

Routine surface water data collections for the purpose of evaluating protected uses

(existing and designated), assessing protected uses, and investigating point and

nonpoint source impacts will be conducted. Products include existing use evaluations,

redesignation recommendations, protected use assessments, and evaluations of point

and nonpoint source discharges or impacts. DEP implements a hybrid approach with

respect to selecting monitoring locations. One component includes a fixed station,

Pennsylvania Water Quality Network (WQN) where stations are established indefinitely

for the purpose of developing long-term datasets that allow for the development of water

quality trends, the ability to measure the effectiveness of water management programs,

the ability to develop water quality assessment methods and standards, and to provide

data that will inform permitted activities. The WQN also includes locations where data is

collected at fixed intervals of five years. Every five years data is evaluated to determine

if the data is serving the previously identified objectives. After each five-year cycle new

stations may be added, stations may be discontinued, and some stations may be

modified.

In addition to the WQN, fixed station approach, more intense short-term locations are

established to evaluate protected uses, assess protected uses and to evaluate point

and nonpoint source impacts. Short-term stations are chosen to ensure that data

representative of conditions in a given stream reach will be obtained. Factors

considered in locating these stations include watershed land uses, volume and chemical

characteristics of known point source wastewater discharges, physiographic and

demographic conditions that contribute to non-point source problems, and stream


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hydrology. Every effort is made to sample representative, homogeneous, low-flow water

columns at comparable locations. However, because weather, stream flow conditions,

and accessibility vary, observations of instream and riparian physical characteristics are

recorded for reference in estimating water quality under various conditions. Data

collection includes biological, chemical and physical data.

Frequency of sampling varies depending on sampling objectives and are described in

Pennsylvania’s Surface Water Quality Monitoring Network Manual (Pulket and

Lookenbill 2020). At WQN stations, waters samples are collected bimonthly (water

samples at ambient and bay loading stations are collected monthly) as discrete samples

at mid-channel, mid-depth on small streams and laterally composited, depth integrated

samples on larger streams or rivers. Semi-quantitative benthic macroinvertebrate

samples are collected annually according to DEP data collection protocols. At WQN

lake stations, an annual qualitative plankton tow is substituted for semi-quantitative

macroinvertebrate sampling. Lake water samples (except Lake Erie) are collected once

a year during summer stratification and consist of a sample collected one meter below

the surface and a sample collected one meter off the bottom at the deepest part of the

lake. Sampling schedules for short-term stations are either limited to one-time

collections and/or governed by protocol and the water quality criteria evaluation needs.

Since data collection is ongoing, only a general timeline is provided below (Table 1).

Sample collection and reporting requirements are further described in Section B of this

QAPP. WQN stations and sampling objectives can be found appended to DEP’s

Monitoring Book as the WQN Manual (Pulket and Lookenbill 2020).

Table 1. General timeline schedule

Month

(represents monthly sequence from project start and not calendar months)

Activity 1 2 3 4 5 6 7 8 9 10 11 12

File Search

Field Recon.

Field Survey

Laboratory Work-up

Report Writing


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Project fiscal information is described as follows:

Data Collection

Office – Planning, file searches, contacts with resource persons

2 workdays

Field – Reconnaissance, water sample collection, field observations 6 workdays

Taxonomic Identifications – Biological samples 4 workdays

Laboratory Support – Chemical/physical/biological analyses 2 workdays

Report Preparation – Data compilation, analysis, and interpretation,

recommendations, draft report preparation, comment review, revisions

10 workdays

TOTAL 24 workdays

A7. Quality Objectives and Criteria

To meet project objectives, biological, physical, chemical or a combination of the three

types of data are collected. Biological data collected for evaluation and assessment

purposes includes macroinvertebrate, fish, fish tissue , mussels, algae/periphyton and

bacteria data. The collection of physical characteristics in the form of habitat evaluations

can also be used to make evaluations and assessments. Stream flow and discharge

can also be collected as supplemental information to make an informed decision.

DEP data collection protocols and assessment methods are part of a larger conceptual

framework that DEP uses to collect quality data and make decisions on various surface

water matters across Pennsylvania. This larger conceptual framework begins with

technical documentation that contains the technical support information and literature to

ensure accurate and precise data analysis and interpretation.

DEP technical documentation can be found at:

https://www.dep.pa.gov/Business/Water/CleanWater/WaterQuality/Pages/Macroinverte

brates.aspx

DEP technical documentation includes characterizations of data quality and quantity to

characterize water quality, biological assemblages, or other parameters with minimal

uncertainty in comparison to relevant WQS. It also includes performance criteria that

data need to achieve in order to make accurate protected use (designated and existing)

evaluations and assessments. General quality control specifications including the

frequency of blanks and duplicates are described throughout this QAPP, and more

specifically in the technical documentation, data collection protocols, and as

appropriate, in the assessment methods. Laboratory quantitation limits can be found in

the Quality Assurance Manual for the PA DEP Bureau of Laboratories (DEP 2019).

Representativeness is addressed in the Monitoring Book Chapter, 2 Sampling Design


E-15

and Planning (Lookenbill 2020) and more specifically, within the appropriate data

collection protocols. Precision estimates for biological methods are described in detail in

respective technical documentation as is detailed development efforts that resulted in

these precision statistics.

Use (designated and existing) evaluations and use assessments based on

physicochemical data can be collected via discrete chemistry data collection or

continuous instream monitoring (CIM). Sediment samples, passive samplers and in-situ

discrete field observations may also be collected to provide additional insight into water

quality conditions. To ensure discrete chemistry samples are collected with the analytes

of interest, the DEP provides specific Standard Analysis Codes (SAC) which collectors

are required to use for chemical and biological samples submitted to DEP Bureau of

Labs. Any deviation from the SACs provided should be approved by the DEP Project

Manager. In addition, at the discretion of WQD, analytes can be added or removed at

any time. Addition of analytes may be warranted if limiting conditions are met and an

analysis to include a more robust suite is needed (e.g. the addition of dissolved

constituents, the addition of phaeophytin, etc.).

All data collection will be conducted according to DEP reviewed and approved

monitoring protocols. Protocols specific to these projects are found in Water Quality

Monitoring Protocols for Streams and Rivers (Monitoring Book, Lookenbill and Whiteash

2021), DEP’s Lake Assessment Protocol (Lathrop 2015), DEP’s Chlorophyl-a Sampling

Method in Lakes, Ponds, and Reservoirs (Lathrop 2017) and DEP’s Evaluations of

Phosphorus Discharges to Lakes, Ponds, and Impoundments (DEP 2013a).

Quality control efforts specific to these projects will be further described in Section B of

this QAPP and are briefly described herein. All biological and chemical samples shall

undergo the collection of replicates every 20 samples or once per project. Chemical

samples shall also undergo collection of blanks every 20 samples or once per project. If

sampling equipment is noted as malfunctioning at time of collection, it should be noted

in a field notebook or on a field data sheet and the DEP project manager should be

informed. If corrective action can be made in the field, the collector should take the

necessary actions to problem-solve. If a solution cannot be determined in the field, field

work should cease and be rescheduled until the problem is solved. For instance, if the

chemistry filtering equipment breaks or malfunctions, no samples should be taken and

sampling should be rescheduled; or for instance, if a field meter probe gives erroneous

readings the collector should determine if calibration drifted, requiring equipment

recalibration. Further information regarding probe accuracies warranting recalibration

are outlined in Hoger et al. 2017; criteria are summarized in Table 2. Additionally,


E-16

assurance of sample integrity is required of any collector. Steps should be taken to

avoid contamination of the sample by an outside source; if at any point during sample

processing contamination is observed, the sample must be properly disposed of and

recollected.

Table 2. Calibration criteria for field measured parameters

Field Parameter Calibration Criteria Warranting Equipment Recalibration


Specific Conductance ± 5 μS/cm or 3%, whichever is

greater

pH ± 0.2 units


Turbidity ± 0.5 FNU or ± 5%, whichever is

greater

Furthermore, chemical samples go through an accuracy check by requiring spikes,

duplicate and blank analyses. Accuracy is determined by routine laboratory protocol

described in the Quality Assurance Manual for the PA Department of Environmental

Protection Bureau of Laboratories (DEP 2019) and in test specific Standard Operating

Procedures (SOP). The BOL Quality Assurance Manual is provided in Supplemental

Document # 1 and SOPs can be made available upon request.

Sensors used for field chemistry measures, whether discrete readings or continuous

deployments, are selected that have a range that will cover expected environmental

conditions for the project, with sufficient precision and accuracy. For all continuous

deployments and preferably for all discrete readings as well, optical DO sensors will be

used as they have higher degrees of accuracy and hold calibration much better. Further

details on sensor selection can be found in the Sensor Selection section of Hoger et al.

(2017).

A8. Training and Certifications

All data collectors need to be properly trained prior to involvement in data collections

and data management. Ensuring data collectors are properly trained is the responsibility

of the quality assurance manager and/or project manager. Training involves reading

DEP’s protocols and training by DEP senior staff or program specialists. New collectors

will go through a training period at the discretion of the supervising biologist or water

program specialist. DEP staff who perform macroinvertebrate identification are required

to maintain Society for Freshwater Science Ephemeroptera Plecoptera Trichoptera

(EPT) East Genus-Level Certification. Any new biologist will endure quality assurance


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audits to ensure adherence to protocols. In addition, an auditor shall accompany an

individual collector on at least one survey at a minimum of every five years. The auditor

shall also randomly select any documentation to verify that data is accurate and

complete, and that appropriate analytical techniques are used. The auditor will maintain

records for each individual to include: (1) the date of the audit; (2) a list of protocols for

which the individual was evaluated; (3) any deficiencies noted; and

(4) recommendations offered and corrective actions taken. Training and quality

assurance records will be maintained in DEP electronic training record files.

All taxonomy should be completed by specialized and/or certified taxonomists.

Taxonomists may be in-house DEP biologist or be contracted out to recognized

laboratories. Taxonomists should use the most updated resources to aid in taxonomic

identifications. Furthermore, taxonomists should undergo QA/QC checks on 10% of

samples identified by a certified taxonomist.

The laboratories used for sample analyses must be accredited. The DEP Bureau of

Laboratories (BOL) will primarily be used for laboratory analyses. BOL is accredited by

the National Environmental Laboratory Accreditation Program (NELAP) through the

New Jersey and Pennsylvania Department of Environmental Protection (DEP)

accreditation bodies. If at any time during the project a new laboratory needs selected to

process samples, the laboratory needs to be a DEP contracted lab and accredited by

bodies within the NELAP.

A9. Documents and Records

Historic and current QAPP documents are housed in WQD’s shared electronic files that

are backed up according to DEP’s information technology security protocols.

Documents are to be maintained and retained permanently. This QAPP will be internally

reviewed annually and will be submitted to EPA for reapproval if material changes are

made. Any amendments or updates to the document should be labeled as such. Every

5 years and as needed, this QAPP should be updated, revised and approved as a new

document that designates signatures and the date of final approval. This will aid in

keeping track of 5-year QAPP resubmission as required by the EPA. Updated versions

will be shared with those on the distribution list given above. Table 3 characterizes

changes to this controlled, QAPP over time. The most recent version is presented in the

top row of the table. Quality assurance data can be obtained through a request to the

DEP Quality Assurance Officer.

Table 3. QAPP document control


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Document

Control

Version #

Title History/Changes Effective

Date

TBD QAPP: Data collection for the

purpose of existing use

evaluations, protected use

assessments, and evaluations

of point and nonpoint source

impacts to surface waters

Previous QAPPs

updated and

consolidated into

Programmatic QAPP

August 1,

2021

391-3200-

002

Fixed Station Surface Water

Quality Network Monitoring

September 22,

2014

384-3200-001

384-3200-001 384-3200-001

Instream Comprehensive

To further describe documents and record keeping, each step of the data collection and

interpretation process is outlined below. Descriptions include creation, maintenance and

retention of records and responsible persons.

Data Collection

Data collected should be recorded on ‘Data Collection Forms’ provided in the

Appendices of DEP’s Monitoring Book (Lookenbill and Whiteash 2021). Additional field

notes can be written on the ‘Data Collection Forms’ or documented in a field notebook.

These ‘Data Collection Forms’ and notes will be used to ascribe in-situ water quality

conditions associated with biological and analytical chemistry sample results.

All benthic macroinvertebrate and fish samples should be given a unique station

identification (ID) number that includes data collected, time collected and collector name

(yyyymmdd-HHMM-Collector). This station ID shall be retained with each sample

through the entire sampling and subsampling processes. Benthic macroinvertebrate

samples and associated data should be entered into DEP’s Streams and Lakes

Integrated Management System (SLIMS) database. Fish tissue data will be entered in

DEP’s Samples Information System (SIS); and, fish, mussel and periphyton community

data is stored in Excel or Access database spreadsheets. For water samples, the field

data, sample and station descriptions, and sample numbers should be entered into SIS

according to Shull, 2017 and retained with the collector until the report is final. DEP

databases reside on enterprise data systems that are backed up and protected

according to DEP information technology security protocols. Record retention for water

quality data is permanent.


E-19

At the end of each sampling season, all ‘Data Collection Forms’ should be filed and

retained for future reference. A filing system will be maintained by each collector. It is

recommended that the collector keep hard copies of completed ‘Data Collection Forms’

as well as maintain specific folders on their computer as an electronic backup labeled

for each data type as a repository for all ‘Data Collection Forms’ (i.e. Macroinvertebrate

Data, Fish Data, CIM Data, WQN Data, etc.). It is further recommended to help facilitate

order that all ‘Data Collection Forms’ be arranged by date and scanned by year to be

placed in a folder that is labeled as ‘YYYY Fish – ‘Field Data Sheets.’ Paper copies can

be archived at the discretion of the field collector and field notebooks should be retained

by the collector.

Calibration logs and calibration equipment checks should be maintained similar to ‘Data

Collection Forms’. It is recommended that calibration logs be kept with each unique

piece of equipment. At the end of the sampling season, the calibration logs shall be

scanned into an electronic file and paper copies archived. Archives are stored in DEP

electronic files that are backed up and protected according to DEP information

technology security protocols. Paper copies can be archived at the discretion of the

equipment manager.

Data Processing

Biological samples that have been processed and identified will be enumerated on

bench sheets found in Appendices of DEP’s Monitoring Book (Lookenbill and Whiteash

2021) Benthic macroinvertebrate taxonomic data will be entered into DEP’s SLIMS

database. Fish, mussel and periphyton community data will be entered into Excel and

Access databases.

All chemical and physicochemical data with the exception of continuous instream

monitoring data are entered into DEP’s SIS database (Shull 2017b). Date, time, location

and the unique sample identifier will be stored in the SIS database. This database will

be used by the BOL to import their analytical results to match field information; BOL will

do this according to the directive found in BOL’s Quality Assurance Manual (DEP 2019).

Each collecting biologist is responsible for following-up on the status of samples

submitted. This includes ensuring samples are analyzed, cross checking samples with

replicate and blank samples and exporting the data from SIS for interpretation.

Continuous instream monitoring data will be housed in a database capable of handling

large continuous files. DEP uses AQUARIUS database and software to store, manage,

and visualize continuous water quality data. AQUARIUS allows for documenting and


E-20

performing corrections of the data according to Continuous Physicochemical Data

Collection Protocol (Hoger et al. 2017).

In preparation for data interpretation and report writing, formatted templates containing

formulas and report requirements are provided to collectors. These templates or

another suitable data organization matrix may be used. If the latter option is used by the

biologist, they must ensure all calculations and visual displays maintain integrity. This

option requires additional quality assurance by a second (or third party if needed) and

the Project Manager before contents can be used to generate reports.

Reports

Data will be reported in the form of tables, figures, maps and/or charts. If warranted, a

full report will be written. Scenarios in which a full report is needed include but is not

limited to redesignation recommendations, integrated report delistings and phosphorus

control recommendations. Redesignation report guidance can be found in DEP’s Water

Quality Antidegradation Implementation Guidance, Document Number 391-0300-002

(DEP 2003). Delisting requirements can be found in DEP’s Assessment Determination

and Delisting Methods (Pulket and Shull 2021). Phosphorus control reports will follow a

general format as described in DEP’s Evaluations of Phosphorus Discharges to Lakes,

Ponds, and Impoundments (DEP 2013a). Continuous instream monitoring reports are

posted on the DEP website at:

https://www.dep.pa.gov/Business/Water/CleanWater/WaterQuality/Pages/CIMReports.a

spx.

Reports are not limited to those items described in the guidance and methods

documents, some projects provide for special cases that require additional analyses

and discussions; these additions can be included at the discretion of the reporting

biologist.

Reports should go through a review process by a second party to address any grammar

or reporting errors/discrepancies. Corrective action should be taken by the report writer

and reviewer; if no agreement can be made, the reviewing party should reach out to the

DEP Project Manager for an additional review. Once finalized, a copy should be dated

and filed appropriately. For final filing the document should be converted from a .docx

file to a .pdf file.

Final Filing

Reports which have been reviewed and finalized should be saved in DEP’s electronic

stream files. Stream files are organized by a unique 5-digit code, which is associated


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with DEP’s historic stream codes associated with the NHD Flowline flowing through the

waterbody, and the stream, river or lake name. Reports shall remain in the stream files

permanently.

Management of QA Record

QA records including training records, quality assurance audits and corrective actions

will be maintained in DEP electronic training record files permanently.


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SECTION B: DATA GENERATION AND ACQUISITION

B1. Experimental Design

Project design will be applied to surface waters statewide. The type of experimental

project design used is based off data of interest. Collectors should use DEP’s

Monitoring Book (Lookenbill and Whiteash 2021) to develop a sampling plan conducive

to each project’s objective. The sampling plan will be objective specific but typically

include a combination of:

• Chemical Data Collection

o In-Situ Field Meter and Transect Data Collection

o Discrete Water Chemistry

o Continuous Water Chemistry

• Physical Data Collection

o Habitat Evaluation

o Water Flow Data Collection

• Biological Data Collection

o Benthic Macroinvertebrates

o Fish

o Bacteria

Site location selection is fluid from year to year based on regional and state priorities. In

addition, environmental factors play a role in selecting sites year to year. For this

reason, site specific maps are not provided in this QAPP. Site locations are documented

as locational attributes in the SLIMS and SIS applications.

Biological samples are collected across a transect or throughout a large portion of the

waterbody to ensure inclusion of all available habitat. In flowing waterbodies water

samples are collected at mid-channel, mid-depth unless stream width, hydrology,

discharge locations or volumes, or observed biological conditions indicate stratification

of flow than integrated type sampling is used. Lake water samples are collected with the

use of a Van Dorn or Kemmerer sampler at a depth of one meter below the surface and

one meter above the lake’s bottom. Parameters frequently analyzed are listed in

Table 3. All water chemistry samples are cooled to less than or equal to 6ºC without

freezing and shipped to the laboratory. Additional parameters may be required based on

the specific nature of the waterbody survey; details regarding field collection

requirements and laboratory test methods can be obtained upon request.


E-23

Table 4. Analyte Tests (commonly used Standard Analysis Code 087)

Test Codes Test Description Test Method

410 ALKALINITY AS CaCO3 @ pH 4.5 SM 2320B 01106H ALUMINUM, DISSOLVED (WATER & WASTE) BY ICPMS EPA 200.8 01105H ALUMINUM, TOTAL (WATER & WASTE) ICPMS EPA 200.8 00608A AMMONIA DISSOLVED AS NITROGEN EPA 350.1 00610A AMMONIA TOTAL AS NITROGEN EPA 350.1 01007A BARIUM, TOTAL (WATER & WASTE) BY ICP EPA 200.7 01022K BORON, TOTAL (WATER & WASTE) BY ICP EPA 200.7 01025H CADMIUM, DISSOLVED (WATER & WASTE) BY ICPMS EPA 200.8 00916A CALCIUM, TOTAL (WATER & WASTE) BY ICP EPA 200.7 01040H COPPER, DISSOLVED (WATER & WASTE) BY ICPMS EPA 200.8 01042H COPPER, TOTAL (WATER & WASTE) BY ICPMS EPA 200.8 00631A DISSOLVE NITRATE & NITRITE NITROGEN EPA 353.2 00671A DISSOLVE ORTHO PHOSPHORUS EPA 365.1 00602A DISSOLVED NITROGEN AS N SM 4500-NC 00666A DISSOLVED PHOSPHORUS AS P EPA 365.1

900 HARDNESS, TOTAL (CALCULATED) SM 2340 B 01046A IRON, DISSOLVED (WATER & WASTE) BY ICP EPA 200.7 01045A IRON, TOTAL (WATER & WASTE) BY ICP EPA 200.7 01049H LEAD, DISSOLVED (WATER & WASTE) BY ICPMS EPA 200.8 01051H LEAD, TOTAL (WATER & WASTE) BY ICPMS EPA 200.8 01130A LITHIUM, DISSOLVED (WATER &WASTE) BY ICP EPA 200.7 01132A LITHIUM, TOTAL (WATER & WASTE) BY ICP EPA 200.7 99020 LOW BROMIDE BY IC EPA 300.1 B

00927A MAGNESIUM, TOTAL (WATER & WASTE) BY ICP EPA 200.7 01056A MANGANESE, DISSOLVED (WATER & WASTE) BY ICP EPA 200.7 01055A MANGANESE, TOTAL (WATER & WASTE) BY ICP EPA 200.7 01065A NICKEL, DISSOLVED (WATER & WASTE) BY ICP EPA 200.7 01067A NICKEL, TOTAL (WATER & WASTE) BY ICP EPA 200.7 82550 OSMOTIC PRESSURE, MOS/KG BOL 3003 403 pH, LAB (ELECTROMETRIC) SM 4500-H+ B

00937A POTASSIUM, TOTAL (WATER & WASTE) BY ICP EPA 200.7 01147H SELENIUM, TOTAL (WATER & WASTE) BY ICPMS EPA 200.8 00929A SODIUM, TOTAL (WATER & WASTE) BY ICP EPA 200.7

95 SPECIFIC CONDUCTIVITY @ 25.0 C SM 2510B 01082A STRONTIUM, TOTAL (WATER & WASTE) BY ICP EPA 200.7 00403T TEMPERATURE AT WHICH PH IS MEASURED SM 4500-H+ B

940 TOTAL CHLORIDE-ION CHROMATOGRAPH EPA 300.0 70300U TOTAL DISSOLVED SOLIDS @ 180C BY USGS-I-1750 USGS I-1750 00630A TOTAL NITRATE & NITRITE NITROGEN EPA 353.2 00600A TOTAL NITROGEN AS N SM 4500-NC

680 TOTAL ORGANIC CARBON SM 5310 C 70507A TOTAL ORTHO PHOSPHORUS AS P EPA 365.1 00665A TOTAL PHOSPHORUS AS P EPA 365.1

945 TOTAL SULFATE-ION CHROMATOGRAPH EPA 300.0 530 TOTAL SUSPENDED SOLIDS USGS I-3765

01090A ZINC, DISSOLVED (WATER & WASTE) BY ICP EPA 200.7 01092A ZINC, TOTAL (WATER & WASTE) BY ICP EPA 200.7


E-24

B2. Sampling Protocols

Biological data collection will be conducted according to protocols described in Chapter

3 of DEP’s Monitoring Book (Lookenbill and Whiteash 2021). Depending on waterbody

classification, benthic macroinvertebrate data collection shall follow one of the four

stream macroinvertebrate data collection protocols (riffle-run, limestone, multihabitat or

semi-wadeable large river streams). Regardless of macroinvertebrate protocols used,

all samples will be processed according to Macroinvertebrate Laboratory Subsampling

and Identification Protocol (Brickner 2020). Fish tissue will be collected according to

DEP’s Fish Tissue Data Collection Protocol (Wertz 2021). Bacteriological data will be

collected according to DEP’s Bacteriological Data Collection Protocol (Miller 2021).

These three types of biological data will be used for the purpose of conducting use

evaluations and use assessments.

Other biological data that may be collected for purposes of describing water quality

conditions and supporting evaluation and assessment decisions are briefly described

herein. Fish community data will be collected according to DEP’s Fish Data Collection

Protocol (Wertz 2021) and mussel community data will be collected according to DEP’s

Mussel Data Collection Protocol (Spear et al. 2017). Periphyton and chlorophyll-a

samples from streams and rivers will be collected according to DEP’s Periphyton Data

Collection Protocol (Butt 2017). Plankton, macrophyte and chlorophyll-a samples from

lakes will be collected according to DEP’s Quantitative Plankton Sampling Method for

Lakes (DEP 2015b), Aquatic Macrophyte Coverage Procedures for Lake Assessments

(DEP 2015a), and Chlorophyll-a Sampling Method for Lakes, Ponds and Reservoirs

(Lathrop 2017) respectively.

Physical data collection in streams and rivers can include stream habitat assessments

and water flow data collection. Descriptions of protocols to collect physical data are

found in Chapter 5 of DEP’s Monitoring Book (Lookenbill and Whiteash 2021). Physical

habitat assessment data collection will be completed when employing all biological data

collection protocols.

Water samples will be collected according to Shull, 2017; modifications may be made

depending on waterbody type and size. All sampling equipment should be rinsed with

surface water three times prior to collection and/or transfer. Most water samples are

collected into HDPE plastic bottles with the exception of organic chemistry samples

which are to be collected in amber glass bottleware. Water samples containing water for

metal analyses should be fixed with HNO3 to a pH of <2. Water samples containing

water for ammonia, phosphorus and organic carbon analyses should be fixed with

H2SO4 to a pH of <2. Samples are immediately placed on ice until submission to BOL.


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Samples should be submitted the day of sampling or the morning after the sampling

event to ensure 48-hour holding times are not exceeded. Analytes with 48-hour holding

times are nitrate, phosphorus and turbidity. Alkalinity has a holding time of 14 days,

specific conductivity, bromide, sulfate and total nitrogen have 28-day holding times; all

other parameters have a holding time of six months or longer. After each use, water

sampling equipment should be thoroughly rinsed with deionized water and cleaned

according to protocols in DEP’s Monitoring Book (Lookenbill and Whiteash 2021). All

other chemistry collections, such as sediment sampling and passive sampler analysis

should follow methodologies found in Chapter 4 of DEP’s Monitoring Book (Lookenbill

and Whiteash 2021).

With any data collection protocol used during a sampling project, it is recommended that

in-situ field meter water quality readings be taken. Measurements could be a single

discrete reading to describe current water quality conditions, transect readings across

the width of the stream or river to verify homogeneity of the water system, or profile

depth reading to determine stratification of larger bodies of water such as

impoundments and lakes. To collect in-situ readings, a calibrated field meter is used. At

a minimum, temperature, pH, dissolved oxygen (DO) and specific conductance

measurements are recorded; other parameters are optional. The field meter should be

calibrated according to procedures outlined in Hoger, 2020 and guidelines referenced

from the equipment’s operation manual. A summary of calibration recommendations is

provided in Table 5. DEP does not use any one specific field meter, to that end specific

guidance on calibration operating procedures cannot be made. At the end of the field

day, calibration checks are recommended as a check against standards to ensure

readings are within the criteria found in Table 2. The collector should also perform

calibration checks anytime field parameters do not make sense for expected conditions.

At the end of the day, the collector should ensure the field meter is free from debris prior

to storage. To clean the field meter equipment, run clean tap water over the probes to

rinse large particles; if a more thorough cleaning is required use a small amount of soap

and a soft bristled brush on the probes. For storage, a small amount of tap water or pH

4 standard should be placed in the storage cup; probes should not be submerged for

overnight or long-term storage.

In addition to discrete physicochemical data, monitoring objectives may require the

collection of continuous physicochemical data according to Hoger et al., 2017. Discrete

physicochemical data typically includes at least four parameters: water temperature,

specific conductance, pH and dissolved oxygen. Continuous water quality monitors

should be calibrated according to procedures outlined in Hoger, 2020 and guidelines

referenced from the equipment’s operation manual. A summary of calibration


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recommendations is provided in Table 5. Calibration follows guidelines referenced from

the equipment’s operation manual. DEP does not use any one specific continuous water

quality monitor, to that end specific guidance on calibration operating procedures cannot

be made. Field meters are used during servicing of a continuous water quality monitor

(1) as a general check of reasonableness of monitor readings, (2) as an independent

check of environmental changes during the service interval, and (3) to make cross-

section surveys or vertical profiles in order to verify the representativeness of the

location of the sonde in the aquatic environment. Data collected by the field meter

should not be used to calibrate the instream monitor. However, some instream monitors

cannot be calibrated, in which case the calibrated field meter must be used in the

calibration correction of monitor records. In addition, DEP may use a calibrated field

meter measurement for undocumented data error corrections (a potential result from the

DEP deployment method, discussed in detail Hoger et al. 2017). Independent field

measurements must be made before, during, and after servicing the monitor in order to

document environmental changes during the service. Side-by-side measurements

between the field meter and the monitor are made by placing the calibrated field meter

as close to the monitor sensors as possible. The field meter should refresh data on the

display at 10 second intervals, or more frequently, to ensure the most accurate data are

recorded during measurements.

B3. Sample Handling and Custody

For biological samples, macroinvertebrates and fishes, samples can be processed for

enumeration by the collector or by other designated staff. Sample containers are

labeled with date, time and user (yyyymm-hhmm-user); stream name; location and

number of containers per sample. A database and archive system shall be maintained

for all samples in which identifications have been completed. Each Regional Office shall

maintain their own archiving system; Central Office will maintain their own archiving

system which can be used by Regional Offices if requested.

Sample handling and chain of custody for chemistry samples should be followed

according to the description provided in Shull 2017a. Steps are summarized herein. The

field collector will use a unique identifier, a seven-digit number that contains a four-digit

collector number and a three-digit sample number. The collector is responsible for

recording locational information and the date and time of the sample. The above

information should be recorded on a sample submission form which is submitted to BOL

at time of relinquishment. Samples can be submitted one of two ways. The collector can

hand deliver samples to BOL’s sample receiving. Under this option, the collector

unloads cooler contents and places samples in designated refrigerators and/or freezers;

the sample submission form remains with the samples. The collector also has the option


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of using a courier service contracted by the DEP. Under this option the collector will

drop coolers at a designated location, located throughout the state, for courier pick-up

and delivery to BOL. The collector should insert their sample submission form into a

plastic sealable bag or whirl-pack (preferably double bagged) and tape it to the inside lid

of the cooler. It is recommended that collectors either scan or take a picture of sample

submission forms for their records. Upon receipt by BOL staff, the temperature of each

sample bottle is measured using a calibrated infrared thermometer to ensure proper

shipping, handling and holding requirements. If samples are found to be above 6°C,

BOL will flag the sample and contact the collector. The flag will remain with the sample

and the reporting biologist shall qualify the data.

Continuous physicochemical data, including operational and maintenance records, are

recorded on portable electronic devices and transferred to AQUARIUS.

Table 5. Summary of Calibration Recommendations

Parameter Calibration Notes

Temperature Check against NIST certified equipment

pH Three-point calibration (4,7,10) Two-point calibration acceptable if conditions are known

Dissolved Oxygen (DO)

100% water-saturated air: • Water in bottom of calibration cup (do not submerge sensor) • Calibration cup on loosely to prevent pressure buildup • Run a wipe cycle, if possible, to remove water droplets on sensor

• Calibration onsite to avoid influence of barometric pressure differences. • Check throughout the day. • Allow plenty of time in cold weather.


One-point calibration with checks • Calibrate at a high value (1000 μS/cm) • Check at a low value (100 μS/cm) • Rinse with Deionized (DI) water and check in air (should be 0 μS/cm)

If values above 1000 μS/cm are expected, also check in a higher standard.


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B4. Analytical Methods

Table 3 above references analytical methods used in analyses of discrete chemistry

samples. The BOL is responsible for setting SOPs for analytical analyses. BOL SOPs

are currently not housed with the WQD but can be obtained upon request. If at any time

during lab analyses sample integrity is compromised, the chemist is responsible for

documenting the flag in the final BOL results report. Occasionally, holding times are

exceeded or samples are incorrectly preserved; if so, BOL notifies the collector

immediately. The collector must then respond to BOL within 10 days to request or

decline results. The collector’s supervisor and any other interested parties must be

copied on their response. If analytes used to make assessment decisions are impacted

by either lab equipment failure or collection error, the sample should be recollected.

While not ideal, the collector may accept the data for the analytes; though, results

should not be used to make assessment determinations.

Other data including biological, physical, field meter and continuous water quality that is

not submitted to BOL are analyzed according to specific data collection protocols and

assessment methods.

B5. Quality Control

Data Collection Audits

All data collected in the field need to be recorded for verification purposes in the

appropriate field data sheets or field notebooks. Field meters used for in-situ water

quality readings should be calibrated according to manufacturer’s instructions at the

start of each field outing. A calibration log should be kept with the field meter and

archived in-house for reference at the end of each sampling year. Calibration checks

should be performed throughout the day if multiple samples will be collected. Results of

calibration and preventive maintenance recommended by the manufacturer must be

recorded in an equipment logbook maintained for each piece of equipment. Dates of

equipment use, calibration results and operator maintenance activities are recorded.

Duplicates for water samples for chemical analysis are collected at least once on each

survey or once every 20 samples. These samples ensure data completeness and

quality assurance of lab analysis techniques. The field investigator will review sample

results and note if target parameters are detected in the blank and flag samples

accordingly.

Duplicate benthic macroinvertebrate samples should be collected within the same reach

as the initial sampling. Habitat, which was sampled previously should not be resampled,

but rather other similar and suitable habitat should be sampled. For semi-wadable


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benthic macroinvertebrate collections of duplicates should be collected as a site

duplicate; this means if it was determined that there were three water quality zones and

three samples across the width of the river need be collected, the replicate should also

contain three samples each in the same reach as the respective water quality zones. In

addition, it is recommended to collect intra-site and inter-site replicates to discriminate

against variation within and between sites in the same stream reach. Fish, mussel, and

plankton samples should undergo, a similar replicate collection schedule of benthic

macroinvertebrate samples. Periphyton sampling QC is detailed more extensively in

Periphyton Data Collection Protocol (Butt 2017).

Quality control of CIM data incorporates multiple field visits throughout the sampling

period. It is recommended that data be downloaded, equipment be cleaned and

checked against calibration standards to describe the accuracy of the continuous

measurements every three to six weeks. QC documentation collected during field visits

will be used to aid in data management prior to making data final. Methods to perform

field visits and correct data are further detailed in DEP’s Continuous Physicochemical

Data Collection Protocol (Hoger et al. 2017).

Results QC

The protocol for validation of chemical data is found in the Quality Assurance Manual for

the PA Department of Environmental Protection Bureau of Laboratories (DEP 2019).

Each collector is also responsible for checking duplicates and field blanks against

samples used for final reporting and calculations. If values are erroneously off from

duplicate and field blanks, those data must by flagged and further investigated. If a

sample is suspected those data should be removed from the final reporting data set.

The field investigator will review sample results to determine if the original sample and

replicate(s) are less than 10% comparatively. For blanks the collector should note if

target parameters are detected in any blank samples. If discrepancies exist, the

collector should flag samples accordingly and leave out of any data interpretation

analyses.

Continuous data that do not meet the thresholds given in the table Maximum Allowable

Limits for Reporting Data (Hoger et al. 2017) are removed from the dataset and will not

be used for assessment or other regulatory decisions. Continuous data that are graded

“Unverified” due to a lack of quality control checks, will be reported but not used for

assessments or regulatory decisions.


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Protocols for biological data validation are found in DEP’s Monitoring Book, Chapter 3

(Lookenbill and Whiteash 2021). Biologist should submit 10% of each type of biological

sample to the Project Manager for taxonomic verification and QA. To accomplish the

10% quality assurance of all samples identified, taxonomists should submit 10% of the

samples they have identified in a calendar year. Documentation of taxonomic QC

results will be stored in WQD, Central Office and samples which were quality assured

will be indicated as so in the SLIMS database. Similarly, fish identifications will be

documented in the associated Excel and Access databases. Furthermore, all data

entries should undergo verification by checking one’s work. During results retrieval and

data interpretation of results, biological data should again be verified against

enumeration forms and database entries.

Subsampling procedures for benthic macroinvertebrate samples and the sequential

identifications will undergo QA/QC. Biologists who are involved with macroinvertebrate

subsampling will need to be properly trained with documentation that tracks sorting error

as is described in the QAPP, Appendix A. Macroinvertebrate identifications will undergo

QC for each biologist involved in taxonomy. The macroinvertebrate identification QC will

be documented according to details provided in Appendix A. Fish identification QC is

also detailed in Appendix A.

The protocol for validation of chemical data is found in the Quality Assurance Manual for

the PA Department of Environmental Protection Bureau of Laboratories (DEP 2019).

Each collector is also responsible for checking duplicates and field blanks against

samples used for final reporting and calculations. If values are erroneously off from

duplicate and field blanks, those data must by flagged and further investigated. If a

sample is suspected those data should be removed from the final reporting data set.

B6. Equipment Maintenance and Inspection

Collectors are expected to care for their own equipment. Equipment should be

inspected prior to each scheduled field outing. If the equipment malfunctions the

collector should trouble shoot or seek assistance from the program specialist. Parts

should be replaced, or back-up equipment should be used.

Field meters, continuous water quality monitoring and flow meters should be inspected

for malfunctions during each use and/or during routine maintenance visits. Probes

should be inspected for damage such as cracks and scrapes. Calibration logs aid in

tracking a probe’s life. Temperature probes are checked against another piece of

equipment. This should be done periodically to ensure proper readings. DO probe

toppers should be replaced every few years as general maintenance; in addition, zero-


E-31

DO checks should be done periodically to determine how well the probe responds to

changing conditions. To make a zero-DO solution dissolve 0.5g of sodium sulfite and 6-

10 crystals of cobalt chloride in 500mL of water, place the probe in solution and

document response time for DO to reach 0.3mg/L. If the response is slow and readings

do not stabilize within 2-3 minutes the probe may need to be replaced. Probes that

measure pH have a short life span and should be replaced at least every year or sooner

if warranted. DEP records pH millivolts in calibration logs to determine the response life

of the pH probe. If pH millivolts drift outside the ranges indicated below, the probe

should be replaced. All other parameters follow specifications found in the equipment’s

operational manual.

• Buffer 7: readings should be -50 to 50 mV, 0 mV is ideal

• Buffer 4: readings should be +165 to +180 mV from the buffer 7 reading

• Buffer 10: readings should be -165 to -180 mV from the buffer 7 reading.

B7. Instrument Calibration

Field meters used to collect in-situ discrete readings, transects and vertical profiles

need calibrated at the start of each sampling day. Continuous water quality monitors are

calibrated and maintained prior to deployment and during routine maintenance visits.

Flow meters are calibrated by the manufacturer every three years. Calibration logs are

kept separate with each unique piece of equipment. See Table 5 for calibration

recommendations. Calibration logs should contain ‘Before’ calibration and ‘After’

calibration readings. Collectors can find calibration log forms in Appendix B of DEP’s

Monitoring Book (Lookenbill and Whiteash). It is recommended that collectors perform

calibration checks on field meters at the end of day that included collecting transect

and/or vertical profile data. Performing end of day checks aid in verifying data are

accurate according to criteria. These checks can be logged in the same logbook using

the same form as the calibration log, where ‘After’ calibration readings are rendered ‘Not

Applicable (NA).’ More detail on field meter use and maintenance can be found in The

Monitoring Book Chapter 4.1, In-Situ Field Meter and Transect Data Collection Protocol,

(Hoger 2020).

B8. Inspection/Acceptance of Supplies and Consumables

All supplies used in data collection should be inspected by the collecting biologist prior

to initial use. Equipment should be checked over at the time of purchase and/or

receiving. Equipment should be returned to the manufacturer if defects are found. In

addition, equipment quality should be inspected prior to and during each use.

Standards used for calibration and sample preservation should be used within

expiration dates and a log of standards used should be maintained. Expired solutions

should be disposed of or recycled properly.


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B9. Non-Direct Measurements

Non-direct measurements associated with projects are tools to help define watershed

and land use characteristics. These characteristics are provided in the final report. To

obtain land use descriptions, DEP often uses USGS Streamstats or ArcGIS software.

These are tools commonly used in the field of hydrology and are acceptable tools to

obtain watershed and land characteristics.

B10. Data Management

Data management should generally follow that which is described herein and DEP data

collection protocols and assessment methods. Data management starts in the field

through the use of field forms and field notebooks. Data contained in field

documentation are entered or transferred into DEP’s SIS, SLIMS, AQUARIUS, Excel or

Access databases as soon as possible. The SIS database is used to store in-situ

discrete field readings and laboratory results. This database organizes, stores and geo-

references data for multiple end uses within DEP. SIS will generate reports which are

emailed to the collector once all chemistry results have been completed by BOL. The

collector is responsible for tracking and filing these BOL auto-generated reports at their

discretion. The final BOL report is used as documentation that samples have been

marked complete by BOL. SIS will also be used by the reporting biologist to obtain

results for organization and interpretation in a final report. The SLIMS database will

generate IBI metric reports for benthic macroinvertebrate samples. In addition, SLIMS

will assist in documenting use assessment determinations for stream segments and

waterbodies.

Data should further be organized into templated spreadsheets for calculations of

indices, models and Chapter 93 WQS criteria violations. These spreadsheets should be

maintained by the biologist responsible for the data. It is recommended that each lake,

stream, river, or basin have its own working folder on the responsible biologist’s

computer, which is separate from the final “Stream File,” where spreadsheets, tables,

charts and draft reports are created. The information in these files should be organized

into a final report, which is placed in the final “Stream File” folder which was mentioned

in the Final Filling Section of A9 of this QAPP.


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SECTION C: ASSESSMENT AND OVERSIGHT

C1. Assessments and Response Actions

Assessments to be made include progress reports which will be provided to EPA mid-

cycle and annually as is required. Additionally, performance assessments will be

provided to collectors and report writers at the time of auditing; and project assessments

will be made on an as needed basis.

On-going QC by all participating parties should have a line of communication open

between the field biologist, the data assessor and the DEP Project Manager. If at any

point during data collection, processing, interpreting and reporting an issue arises, the

responsible party should reach out to resources by providing a summary of the issue to

include the problem description and the corrective action needed. The responsible party

needs to document steps taken until the issue has been resolved; documentation can

include retaining email chains or creating a timeline spreadsheet.

C2. Reports to Management

Reports regarding ongoing performance and system audits, data quality evaluations

and significant quality assurance problems will be included in the mid-cycle and annual

grant status report to EPA and will be provided to the DEP’s Quality Systems Program

Manager and the WQD, Monitoring Section.

Based on priorities, the field biologist should complete a final report to the DEP Project

Manager in a timely fashion. It is understood that project priorities may change within

the DEP due to time sensitive deadlines; if an extension or assistance in data

management is needed, biologists can request assistance from WQD, Central Office

staff.


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SECTION D: DATA VALIDATION AND USABILITY

D1. Data Review, Verification and Validation

The protocol for validation of biological data is found in DEP’s Monitoring Book, Chapter

3 (Lookenbill and Whiteash). The protocol for validation of chemical data is found in the

Bureau of Laboratories, Quality Assurance Manual for the PA Department of

Environmental Protection Bureau of Laboratories (DEP 2019) and BOL’s SOPs. The

protocol for validation of physicochemical data is found in DEP’s Continuous

Physicochemical Data Collection Protocol (Hoger et al. 2017).

A log is maintained of field instrumentation calibrations, performance, and repairs.

Additionally, the biologist responsible for interpreting data will provide a review of results

to include duplicate and blank comparison and best professional judgement

determinations based on knowledge of the site and equipment used. If detected,

erroneous data will be flagged, described and ultimately removed from the final dataset.

Calculations using indices, models and Chapter 93 WQS criteria are checked during

internal review. All continuous data undergoes final review approval according to DEP

data collection protocols.

D2. Verification and Validation of Protocols

Errors are detected through verification of data by the biologist in charge of the survey,

in-house review of reports, and supporting calculations and field audits. When

problems are noted, the responsible individual is notified, provided with the appropriate

protocol and training, and reevaluated before performing the task again. The auditor

shall maintain the records of any corrective actions on additional auditing forms.

With any data management, integrity is of upmost importance to provide quality results

and analyses of the data. It is necessary that collectors cross-validate all field notes with

data entries into DEP’s data management databases. DEP currently maintains a

biological (SLIMS), discrete physicochemical (SIS) and continuous physicochemical

(AQUARIUS) databases. The collector should quality assure their data at the end of the

sampling event and again prior to the development of any reports. In addition, all reports

will need to be reviewed internally and by the DEP Project Manager prior to submittal to

EPA and prior to distribution to the public to ensure accuracy of the reporting. In

addition, all calculations used to generate index scores, models, and water quality

criteria exceedances will be reviewed for integrity prior to distributing for use by

biologists responsible for interpreting the data. If any problems arise with calculations,

formulas, spreadsheets, databases, etc., which are provided to facilitate data

interpretation, the user should note issues and inform the DEP Project Manager of


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discrepancies and problems, at which time the appropriate corrective action will be

determined, and updated materials will be distributed.

D3. Reconciliation with User Requirements

The data generated from the surveys and reports will be analyzed and reconciled by

cross referencing DEP assessment methods (Shull and Whiteash 2021); assessments

are made in accordance with Section 96.3 and 96.5(b) of DEP’s regulations. The end

product should reconcile with requirements documented in DEP’s Monitoring Book

(Lookenbill and Whiteash 2021), DEP’s Assessment methodology for Streams and

Rivers (Shull and Whiteash 2021), DEP’s Water Quality Antidegradation Implementation

Guidance (DEP 2013b) and DEP’s lake assessment methods which are listed on DEP’s

website

https://www.dep.pa.gov/Business/Water/CleanWater/WaterQuality/Pages/Assessment-

Methodology.aspx.

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LITERATURE CITED

Brickner, M. (editor) 2020. Macroinvertebrate laboratory subsampling and identification

protocol. Chapter 3.6, pages 33–43 in M. J. Lookenbill, and R. Whiteash



Butt, J. S. 2017. Periphyton data collection protocol. Chapter 3.10, pages 88–142 in M.

J. Lookenbill, and R. Whiteash (editors). Water quality monitoring protocols for



DEP. 2019. Quality assurance manual for the Pennsylvania Department of

Environmental Protection Bureau of Laboratories. Supporting Document #1.

DEP. 2015a. Aquatic macrophyte coverage procedures for lake assessments.


Pennsylvania.

DEP. 2015b. Quantitative plankton sampling method for lakes (Modified Standard

Method 1002A). Pennsylvania Department of Environmental Protection,


DEP. 2013a. Evaluations of phosphorus discharges to lakes, ponds, and

impoundments. Pennsylvania Department of Environmental Protection,


DEP. 2013b. Water quality antidegradation implementation guidance, Document

Number 391-0300-002. Pennsylvania Department of Environmental Protection,


Hirsch, R. M., D. L. Moyer, and S. A. Archfield. 2010. Weighted regressions on time,

discharge, and season (WRTDS), with an application to Chesapeake Bay river

inputs. JAWRA 46(5):857–880.









Pennsylvania.

Lathrop, B. F. 2017. Chlorophyl a sampling method in lakes, ponds, and reservoirs.


Pennsylvania. (Available online at:

E-37

https://www.dep.pa.gov/Business/Water/CleanWater/WaterQuality/Pages/Assess

ment-Methodology.aspx.)

Lathrop, B. F. 2015. Lake assessment protocol. Pennsylvania Department of

Environmental Protection, Harrisburg, Pennsylvania. (Available online at:

https://www.dep.pa.gov/Business/Water/CleanWater/WaterQuality/Pages/Assess

ment-Methodology.aspx.)




Miller, S. (editor). 2021. Bacteriological data collection protocol. Chapter 3.11, pages




Pulket, M., and M. J. Lookenbill. Pennsylvania’s surface water quality monitoring

network manual. Appendix D pages 1–71 in M. J. Lookenbill, and R. Whiteash



Spear, R., D. Counahan and Shull, D. R. 2017. Mussel data collection protocol. Chapter

3.9, pages 74–87 in M. J. Lookenbill, and R. Whiteash (editors). Water quality







Shull, D. R. 2017b. Sample information system (SIS) data entry protocol. Chapter 4.6,




Shull, D. R., and R. Whiteash (editors). 2021. Assessment methodology for streams

and rivers. Pennsylvania Department of Environmental Protection, Harrisburg,

Pennsylvania.

Shull, D. R., and M. Pulket. 2021. Assessment determination and delisting methods.

Chapter 5, pages 1–15 In Shull, D. R. and, R. Whiteash (editors). Assessment

methodology for streams and rivers. Pennsylvania Department of Environmental


Wertz, T. A. 2021. Fish data collection protocol. Chapter 3.7, pages 44–63 in M. J.

Lookenbill, and R. Whiteash (editors). Water quality monitoring protocols for



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Wertz, T. A. (editor). 2021. Fish tissue data collection protocol. Chapter 3.8, pages 64–

73 in M. J. Lookenbill, and R. Whiteash (editors). Water quality monitoring



E-39

E.1 BENTHIC MACROINVERTEBRATE SUBSAMPLING AND TAXONOMIC IDENTIFICATION QUALITY CONTROL

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INTRODUCTION

The Department of Environmental Protection, Bureau of Clean Water, Water Quality

Division (WQD) maintains that all biologist involved in the processing and identification

of macroinvertebrate samples be properly trained and audited by WQD staff according

to protocols found in Chapter 3 of DEP’s Water Quality Monitoring Protocols for

Streams and Rivers (Lookenbill and Whiteash 2021).

Steps to ensure the proper identification of benthic macroinvertebrate and fish

taxonomy are summarized as follows:

1) At least 10% of all samples identified by a biologist are to be quality assured by a

certified taxonomist. Identification results between the biologist and certified

taxonomist must be 90% or greater.

2) DEP, at its discretion, may request sample vials for any results entered into

internal databases for QA purposes.

As part of the taxonomic quality control (QC), documentation of quality assurance of

subsampling and identification procedures shall follow that described below.

QC – BENTHIC MACROINVERTEBRATE SUBSAMPLING

Subsampling macroinvertebrates in the laboratory should follow a training, audit and QA

schedule, as described herein. Each new sorter will go through a period of training. This

training requires samples be sorted and checked by supervising staff. A sample here is

defined as being a sample that has been picked to completeness, following the

procedures outlined in the Macroinvertebrate Laboratory Subsampling and Identification

Protocol (Brickner 2020). The individual must follow protocol sorting grid by grid by

taking contents out of the randomly selected grid and placing the contents into a viewing

pan. The sorter will then pick out all organisms one by one and place them in a petri

dish. After each grid is sorted and no remaining organisms can be found, the viewing

pan should be checked by supervising staff. In addition, the supervising staff should

check the petri dish for any specimens that do not belong; specimens that do not belong

include pupae, half body specimens, adult specimens (except aquatic adult

Coleopterans), exoskeletons (exuviae) and specimens degraded beyond identification.

The supervising staff and sorter will track the number of misses per viewing pan and

number of specimens that do not belong after each petri dish check. If the supervising

staff finds the sorter missing more than 10 specimens on their first scan check, the

sorter will be given a second opportunity to re-look with instruction given to pay better

E-41

attention to commonly missed specimens. A re-look does not count against the sorter

and their training; however, the supervising staff should document a re-look was

required and should document the instruction given. During the initial training, if grid

error (Equation 1) calculated across all grids sorted during the full sample sort is greater

than 10% the individual fails their training and will need to go through a subsequent

training period until the individual passes; it is recommended that a minimum of three

full samples be checked prior to completing an audit for sign-off of independent sorting,

but this is left up to the discretion of supervising staff.

Once training is passed the individual will complete one more ‘full’ sample sort for the

final audit. If the four or more grids sorted out of Pan 1 results in a GE of greater than

5%, the audit fails, otherwise the sorter passes and can be cleared to pick without any

oversite. At the discretion of supervising staff, seasonal staff will be audited as needed;

and, agency wide, permeant staff will be audited every three years. An audit will consist

of one sample that is picked to completeness and checked by supervising staff. To pass

the audit, the one sample QA check should yield no greater than 5% grid error

(Equation 1). If the audit fails, the individual will need monitored until a passing grade is

achieved. With two subsequent fails the individual will have to be re-trained. If the audit

passes the individual may continue to sort independently, until the next auditing event.

Equation 1: Grid Error (GE), expressed as a percentage

Described as the mean weighted grid error for the full sample sort. Calculated according

to percent error documented in each pan and weighted by the ratio of organisms per

viewing pan divided by the total number of organisms sorted.

GE = (∑ 𝑠𝑜𝑟𝑡𝐸 ∗ 𝑊𝑖 )/P

Where: sortE = sorting error calculated by number of misses divided by the total number

of

organisms sorted out of the viewing pan; W = weight given to each viewing pan based

on the total population of the full sample sort; P= number of pans sorted

Example:

Pan 1: Number sorted 33, misses 5, petri remove 0, total 38




Total number after four girds 240

Sorting error calculation (sortE):

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Pan 1 = (1-(33/38))*100 = 13.16%

Pan 2 = (1-(41/45))*100 = 8.88%

Pan 3 = (1-(79/90))*100 = 12.22%

Pan 4 = (1-(62/67))*100 = 7.46%

Weight calculation (W):

Pan 1 = 38/240 = 0.158

Pan 2 = 45/240 = 0.188

Pan 3 = 90/240 = 0.375

Pan 4 = 67/240 = 0.279

GE = ((13.16*0.158)+(8.88*0.188)+(12.22*0.375)+(7.46+0.279))/4 = 2.6%

QC – BENTHIC MACROINVERTEBRATE IDENTIFICATION

As described above, biologists are to uphold standards of identifying organisms to

specific taxonomic levels. Ephemeroptera, Plecoptera, and Trichoptera (EPT) taxa are

to be identified to Genus; midges, snails, clams and mussels are to be identified to

Family level; flatworms, proboscis worms, roundworms and moss animalcules are to be

identified to Phylum level; aquatic earthworms and tubificid worms are to be identified to

Class; and water mites are to be identified to Subclass. If for any reason the highest

taxonomic identification requirements cannot be made, the Monitoring Book states that

“…the biologist must forward identifications to a certified taxonomist for further

evaluation…(Lookenbill and Whiteash 2021).” It was noted that 10% of all samples

identified by a biologist are to be QAed by a certified taxonomist. To accomplish the

10% quality assurance of all samples identified, taxonomists should submit 10% of the

samples they have identified for a calendar year. The easiest way to accomplish this is

to flag every tenth sample identified to be submitted for QA. This subset should

represent an even distribution of all samples collected and/or identified for a calendar

year. Samples should be delivered to the DEP staff responsible for sample QA

beginning January first of the succeeding year and no later than February first. DEP

staff, Regional and Central Office, have the option to request the QA of samples that

were not collected or identified by them if they have a specific interest in it (i.e.

permitting).

Samples submitted for taxonomic verification will undergo calculations to determine

percent disagreements in taxonomy, enumeration and Index of Biotic Integrity (IBI)

metrics between the biologist and quality assurer results. Errors documented by the

taxonomic verification QA procedure were developed similarly to that described in

Stribling et al. 2008. Of interest is Percent Taxonomic Disagreement (PTD), Percent

E-43

Enumeration Disagreement (PED), Percent IBI Disagreement (PID). Percent taxonomic

disagreement takes into account differences in specimen identifications between the

biologist and the quality assurer. Individual taxon agreements are determined by

comparing lists, and a percent difference is calculated according to Equation 2;

calculated PTD error should be no greater than 10%. Percent enumeration

disagreement is a calculation that determines the counting error. PED is calculated

according to Equation 3 and should be no greater than 5%. Percent IBI disagreement

considers differences in the IBI scores generated by the two taxa lists (Equation 4);

calculated PID error should be no greater than 10%. If any calculated error, (PTD, PED

or PID) is greater than the 10% or 5% criteria, corrective action be taken. Corrective

action could include an opportunity for the biologist to re-look at the samples, a

conversation between the biologist and quality assurer or the recommendation to seek

further training in the identification of problem taxa.

Equation 2 – Percent Taxonomic Disagreement (PTD), expressed as a percentage

PTD = (1 − [a

Nmax] ) x 100

Where: a = total number of agreements (summed across all individuals and taxa); Nmax

= total number of individuals identified (the greater of the two totals)

Equation 3 – Percent Enumeration Disagreement (PED), expressed as a

percentage

PED = ([(ni−nq)

(ni+nq)]) x 100

Where: ni = number of individuals counted by the biologist; nq = number of individuals

counted by the quality assurer

Equation 4 – Percent IBI Disagreement (PID), expressed as a percentage

PID = ([(Si−Sq)

Sq]) x 100

Where: Si = the IBI score generated by the taxa list provided by the biologist; Sq = the

IBI score generated by the taxa list provided by the quality assurer

QC – FISH IDENTIFICATION

Fish samples identified by DEP biologists should undergo QA/QC similarly to benthic

macroinvertebrate samples. 10% of all samples identified by a biologist are to be QAed

by a certified taxonomist. To accomplish the 10% quality assurance of all samples

identified, taxonomists should submit 10% of the samples they have identified for a

E-44

calendar year. The easiest way to accomplish this is to flag every tenth sample

identified to be submitted for QA. This subset should represent an even distribution of

all samples collected and/or identified for a calendar year. Samples should be delivered

to the DEP staff responsible for sample QA beginning January first of the succeeding

year and no later than February first. DEP staff, regional and central office, have the

option to request the QA of samples that were not collected or identified by them if they

have a specific interest in it (i.e. permitting).

Samples submitted for taxonomic verification will undergo calculations to determine

percent disagreements in taxonomy and enumeration between the biologist and quality

assurer results. Errors documented by the taxonomic verification QA procedure were

developed similarly to that described in Stribling et al. 2008. Of interest is Percent

Taxonomic Disagreement (PTD) and Percent Enumeration Disagreement (PED).

Percent taxonomic disagreement takes into account differences in specimen

identifications between the biologist and the quality assurer. Individual taxon

agreements are determined by comparing lists, and a percent difference is calculated

according to Equation 2; calculated PTD error should be no greater than 10%. Percent

enumeration disagreement is a calculation that determines the counting error. PED is

calculated according to Equation 3 and should be no greater than 5%. If any calculated

error, (PTD or PED) is greater than the 10% or 5% criteria, corrective action be taken.

Corrective action could include an opportunity for the biologist to re-look at the samples,

a conversation between the biologist and quality assurer or the recommendation to seek

further training in the identification of problem taxa.

LITERATURE CITED






Pennsylvania.

Stribling J. B., K. L. Pavlik, S. M. Holdsworth, and E. W. Leppo. 2008. Data quality,

performance, and uncertainty in taxonomic identification for biological

assessments. Journal of North American Benthological Society 27(4):906–919.

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